Bancroft s Theory and Practice of Histological Techniques, International Edition
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836 pages

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This is a brand new edition of the leading reference work on histological techniques. It is an essential and invaluable resource suited to all those involved with histological preparations and applications, from the student to the highly experienced laboratory professional. This is a one stop reference book that the trainee histotechnologist can purchase at the beginning of his career and which will remain valuable to him as he increasingly gains experience in daily practice.

Thoroughly revised and up-dated edition of the standard reference work in histotechnology that successfully integrates both theory and practice.Provides a single comprehensive resource on the tried and tested investigative techniques as well as coverage of the latest technical developments.

Over 30 international expert contributors all of whom are involved in teaching, research and practice.Provides authoritative guidance on principles and practice of fixation and staining.

Extensive use of summary tables, charts and boxes.Information is well set out and easy to retrieve.

Six useful appendices included (SI units, solution preparation, specimen mounting, solubility). Provides practical information on measurements, preparation solutions that are used in daily laboratory practice.

Color photomicrographs used extensively throughout. Better replicates the actual appearance of the specimen under the microscope.

Brand new co-editors.

New material on immunohistochemical and molecular diagnostic techniques.Enables user to keep abreast of latest advances in the field.


Operating microscope
Renal biopsy
Journal of Clinical Pathology
Parkinson's disease
Ammonium molybdate
Hydrochloric acid
Acetic acid
Isotype (immunology)
Potassium acetate
Manganese heptoxide
Reticular fiber
In situ hybridization
Mineral acid
Fluorescent in situ hybridization
Sodium acetate
Protein S
Carcinoma in situ
Polyclonal antibodies
Derivative (disambiguation)
Congo red
Biological agent
Immunoglobulin E
Bottled water
Image analysis
Genetic testing
Tetralogy of Fallot
Tap water
Further education
Connective tissue
Risk assessment
Transmission electron microscopy
Iron deficiency
Distilled water
Tissue (biology)
Ammonium nitrate
Sodium chloride
Organic acid
Multiple sclerosis
Strong acid
Diabetes mellitus
United Kingdom
Scanning electron microscope
International System of Units
Risk management
Nucleic acid
Hydrogen bond
Fatty acid
Electron microscope


Publié par
Date de parution 01 octobre 2012
Nombre de lectures 1
EAN13 9780702050329
Langue English
Poids de l'ouvrage 3 Mo

Informations légales : prix de location à la page 0,0589€. Cette information est donnée uniquement à titre indicatif conformément à la législation en vigueur.


Bancroft’s Theory and Practice of Histological Techniques
Seventh Edition

S. Kim Suvarna
Consultant Pathologist, Histopathology Department, Northern General Hospital, Sheffield, UK

Christopher Layton
Specialist Section Lead in Specimen Dissection, Histopathology Department, Northern General Hospital, Sheffield, UK

John D. Bancroft
Formerly Pathology Directorate Manager and Business Manager, Queen’s Medical Centre, Nottingham, UK
Churchill Livingstone
Table of Contents
Instructions for online access
Cover image
Title page
Preface to the seventh edition
Preface to the first edition
List of contributors
Chapter 1: Managing the laboratory
Chapter 2: Safety and ergonomics in the laboratory
Risk management
Control of chemicals hazardous to health and the environment
Control of biological substances hazardous to health and the environment
Control of physical hazards
Hazards and handling of common histological chemicals
Chapter 3: Light microscopy
Light and its properties
Image quality
The components of a microscope
Magnification and illumination
Phase contrast microscopy
Interference microscopy
Polarized light microscopy
Fluorescence microscopy
Use of the microscope
Setting up the microscope
Chapter 4: Fixation of tissues
Types of fixation
Physical methods of fixation
Chemical fixation
Special fixatives
Compound fixatives
Factors affecting the quality of fixation
Selecting or avoiding specific fixatives
Fixation for selected individual tissues
Useful formulae for fixatives
For metabolic bone disease
Fixation and decalcifation
Fixation for fatty tissue
Chapter 5: The gross room/surgical cut-up
Safety first and last
Specimen reception
Surgical cut-up/specimen dissection/grossing
Thinking before dissection
Specimen dissection plans
Chapter 6: Tissue processing
Tissue microarray
Chapter 7: Microtomy: Paraffin and frozen
Paraffin section cutting
Frozen and related sections
Uses of frozen sections
Cryostat sectioning
Freeze drying and freeze substitution
Frozen section substitution
Chapter 8: Plastic embedding for light microscopy
Ultrastructural studies
Hard tissues and implants
High-resolution light microscopy
Plastic embedding media
Applications of acrylic sections
In situ hybridization
Acrylic plastic processing schedules
Future of acrylic plastic embedding
Chapter 9: How histological stains work
A general theory of staining
Some dyestuff properties
Problem avoidance and troubleshooting
Chapter 10: The hematoxylins and eosin
Alum hematoxylins
Routine staining procedures using alum hematoxylins
Iron hematoxylins
Tungsten hematoxylins
Molybdenum hematoxylins
Lead hematoxylins
Hematoxylin without a mordant
Quality control in routine H&E staining
Difficult sections
Chapter 11: Connective and mesenchymal tissues with their stains
Formed or fibrous intercellular substances
Methenamine silver microwave method
Connective tissue cells
Connective tissues
Connective tissue stains
Chapter 12: Carbohydrates
Classification of carbohydrates
Connective tissue glycoconjugates – the proteoglycans
Other glycoproteins
Techniques for the demonstration of carbohydrates
Lectins and immunohistochemistry
Enzymatic digestion techniques
Chemical modification and blocking techniques
Chapter 13: Pigments and minerals
Endogenous pigments
Artifact pigments
Exogenous pigments and minerals
Chapter 14: Amyloid
Metachromatic techniques for amyloid
Polarizing microscopy
Acquired fluorescence methods
Miscellaneous methods
Fibril extraction
Immunohistochemistry for amyloid
Laser microdissection-proteomics: a new tool for typing amyloid
Evaluation of methods
The future
Chapter 15: Microorganisms
General principles of detection and identification
The Gram stain
Techniques for mycobacteria
Some important bacteria
Fungal infections
A selection of the more important fungi and actinomycetes
The demonstration of rickettsia
The detection and identification of viruses
Viral infections
Prion disease
The demonstration of protozoa and other organisms
Chapter 16: Bone
Normal bone
Techniques for analyzing bone
Processing decalcified bone
Preparation of mineralized bone
Morphometry of bone
Chapter 17: Techniques in neuropathology
The components of the normal nervous system
Techniques for staining neurons
The neuroglia
Neuropathology laboratory specimen handling
Chapter 18: Immunohistochemical techniques
Immunohistochemistry theory
Immunohistochemical methods
Immunohistochemistry in practice
Chapter 19: Immunofluorescent techniques
Preservation of substrate antigens
Primary antibodies and conjugates
Staining procedure
Quality control
Diagnostic histopathology
Chapter 20: Immunohistochemistry quality control
Factors affecting stain quality
Monitoring stain quality
Chapter 21: Molecular pathology
Common reagents
Probes and their choice
Probe preparation and labeling
Preparation of the dilution series
Commercially made probes
Sample preparation
Treatment of solutions and glassware to destroy nuclease activity
Genetic testing: fluorescence in situ hybridization (FISH)
General FISH procedure
FISH set-up
Specific FISH procedure: HER2 FISH (PathVysionTM)
General scoring analysis criteria for FISH
Troubleshooting FISH
Validation of FISH probes in the clinical laboratory
FISH nomenclature
Glossary and definitions of the terminology used in this chapter and in ISH techniques
Chapter 22: Transmission electron microscopy
Tissue preparation for transmission electron microscopy
Specimen handling
Wash buffer and staining
Epoxy resins
Acrylic resins
Tissue processing schedules
Procedures for other tissue samples
Diagnostic applications
Renal disease
Malignant tumors
Non-neoplastic diseases
Chapter 23: Quantitative data from microscopic specimens
Traditional approaches
Image analysis
Image analysis processes
Image analysis software
Specimen analysis
Specimen preparation for image analysis
Multispectral imaging
Appendices: Diagnostic Appendices
Appendix I: Classical histochemical methods
Appendix II: Applications of immunohistochemistry
Appendices: Technical Appendices
Appendix III: Measurement units
Appendix IV: Preparation of solutions
Appendix V: Buffer solutions
Appendix VI: Solubility of some common reagents and dyes
Appendix VII: Mounting media and slide coatings
Appendix VIII: Molecular pathology reagents
Staining methods index
Subject index

is an imprint of Elsevier Limited
© 2013, Elsevier Limited. All rights reserved.
First edition 1977
Second edition 1982
Third edition 1990
Fourth edition 1996
Fifth edition 2002
Sixth edition 2008
The right of Dr. S. Kim Suvarna, Dr. Christopher Layton and Mr. John D. Bancroft to be identified as author of this work has been asserted by them in accordance with the Copyright, Designs and Patents Act 1988.
No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: .
This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein).

Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary.
Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility.
With respect to any drug or pharmaceutical products identified, readers are advised to check the most current information provided (i) on procedures featured or (ii) by the manufacturer of each product to be administered, to verify the recommended dose or formula, the method and duration of administration, and contraindications. It is the responsibility of practitioners, relying on their own experience and knowledge of their patients, to make diagnoses, to determine dosages and the best treatment for each individual patient, and to take all appropriate safety precautions.
To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein.
Churchill Livingstone
British Library Cataloguing in Publication Data
Bancroft’s theory and practice of histological techniques.
 – 7th ed.
 1. Histology, Pathological – Technique.
 I. Theory and practice of histological techniques
 II. Suvarna, Kim. III. Layton, Christopher. IV. Bancroft,
 John D.
 616′.07583 – dc23
ISBN-13: 9780702042263
ISBN: 978-0-7020-4226-3
Ebook ISBN: 978-0-7020-5032-9
Printed in China
Last digit is the print number: 9 8 7 6 5 4 3 2 1
You are familiar with earlier editions of Theory and Practice of Histological Techniques . So, you may be wondering ‘What’s all this about? Why the name change?’ As the only author contributing to this new edition who also contributed to the first edition, and as someone originally recruited by the eponymous John Bancroft and his then co-editor Alan Stevens, it falls to me to offer an explanation. It is simple enough. John has now pulled back into a more hands-off editorial role. Yet his energy and persistence over many years are the key to this publishing epic – seven editions of a technical manual, continuously in print for 35 years, wow! So Churchill Livingstone Elsevier, the publishers, wish to celebrate John’s part in the success of this world-renowned text from its origins to this new edition, both as editor and contributor. Moreover, the successive editions of this remarkable book were, for much of the time, produced in parallel with and enriched by John’s contributions to research and teaching in our field, and of course whilst managing a histopathology department in a large teaching hospital. So let us salute Bancroft’s Theory and Practice of Histological Techniques !

Richard Horobin
Preface to the seventh edition
In the 35 years since the first edition of this book, the histological laboratory has changed dramatically. Whilst some techniques of tissue selection, fixation and section production have remained relatively static, there have been great advances in terms of immunological and molecular diagnostic methodology. Immunohistochemistry and immunofluorescence now have well-defined roles with quality assurance realities, and are to be found throughout the world with pivotal interactions with tissue diagnosis and patient management. In the last 20 years, the progressive development of molecular techniques revolving around DNA and in situ hybridization has permitted the creation of new genetic tests and diagnostic opportunities for the laboratory. These are currently at the forefront of guiding treatment choices for patients. At the same time, these have permitted review of some classic histological tests resulting in a reduced histochemical repertoire in many laboratories. Knowledge of both the old and new is required by trained, as well as trainee, histotechnologists working alongside the pathologist. A thorough grounding in all these aspects of diagnostic methodology is still required.
In producing this edition we were faced with choices about classical and rarely used methodologies, and concluded that many needed to be removed from the text or reduced in volume into the appendices. These include the chapters on lipids, proteins and nucleic acids, neuroendocrine system and cytoplasmic granules and enzyme histochemistry. This has allowed for expansion and update in some areas, particularly the newer diagnostic methodologies. We recognized that some sections on classic stains have not changed dramatically, and have simply reviewed these to ensure that modern relevance has been achieved. Other chapters have been amalgamated, such as the in situ hybridization and genetic testing sections.
There are a number of new contributors for this edition. They include Louise Dunk, who contributed the management chapter and Anthony Rhodes, who updated fixation of tissues. The gross room/surgical cut-up chapter has been rewritten by Kim Suvarna and Christopher Layton. The pigment and minerals chapter has been revamped by Guy Orchard and the amyloid chapter by Janet Gilbertson and Tony Hunt. Neuropathology has been rewritten by Robin Highly and Nicky Sullivan. Some immunohistochemistry and immunofluorescent techniques have required a rewrite reflecting current modalities, and this has been accomplished by Tracy Sanderson, Greg Zardin and Graeme Wild.
Having said this, we are conscious that we are all part of the lineage of previous authors that have contributed to the first six editions of the book. We salute and thank them for their previous work. Indeed, their contribution to the success of this ongoing text cannot be underestimated. We would not wish to single out any one person, or group of individuals, but rather express great thanks to all the previous contributors over the decades that this book has been in existence.
Ultimately, we hope that we have produced a modern and relevant histotechnology text that will be of use to those in training as well as established practitioners across the world. As always, we recognize that this edition is but one step of the ongoing story and hope that colleagues across the world will enjoy and approve of the changes that have taken place.
S. Kim Suvarna, Christopher Layton and John D. Bancroft
February 2012
Preface to the first edition
In recent years histological techniques have become increasingly sophisticated, incorporating a whole variety of specialties, and there has been a corresponding dramatic rise in the level and breadth of knowledge demanded by the examiner of trainees in histology and histopathology technology.
We believe that the time has arrived when no single author can produce a comprehensive book on histology technique sufficiently authoritative in the many differing fields of knowledge with which the technologist must be familiar. Many books exist which are solely devoted to one particular facet such as electron microscopy or autoradiography, and the dedicated technologist will, of course, read these in the process of self-education. Nevertheless the need has arisen for a book which covers the entire spectrum of histology technology, from the principles of tissue fixation and the production of paraffin sections to the more esoteric level of the principles of scanning electron microscopy. It has been our aim then, to produce a book which the trainee technologist can purchase at the beginning of his career and which will remain valuable to him as he rises on the ladder of experience and seniority.
The book has been designed as a comprehensive reference work for those preparing for examinations in histopathology, both in Britain and elsewhere. Although the content is particularly suitable for students working towards the Special Examination in Histopathology of the Institute of Medical Laboratory Sciences, the level is such that more advanced students, along with research workers, histologists, and pathologists, will find the book beneficial. To achieve this we have gathered a team of expert contributors, many of whom have written specialized books or articles on their own subject; most are intimately involved in the teaching of histology and some are examiners in the HNC and Special Examination in Histopathology. The medically qualified contributors are also involved in technician education.
All contributors have taken care to give, where applicable, the theoretical basis of the techniques, for we believe that the standard of their education has risen so remarkably in recent years that the time is surely coming when medical laboratory technicians will be renamed ‘medical laboratory scientists’; we hope that the increase in ‘scientific’ content in parts of this book will assist in this essential transformation.

John D. Bancroft

Alan Stevens
Nottingham, 1977
List of contributors

Caroline Astbury, PhD FACMG
Department of Pathology and Laboratory Medicine Nationwide Children’s Hospital Columbus, OH, USA

John D. Bancroft
Formerly Pathology Directorate Manager and Business Manager Queen’s Medical Centre Nottingham, UK

Jeanine H. Bartlett, BS HT (ASCP), QIHC
Biologist Centers for Disease Control and Prevention Infectious Diseases Pathology Branch Division of High-Consequence Pathogens and Pathology National Center for Emerging and Zoonotic Infectious Diseases Atlanta, GA, USA

David Blythe, FIBMS
Chief Biomedical Scientist HMDS Laboratory Leeds Teaching Hospitals NHS Trust Leeds, UK

Louise Dunk, MSc FIBMS
Lead Laboratory Manager Histopathology Sheffield Teaching Hospitals Sheffield, UK

Alton D. Floyd, PhD
ImagePath Systems Inc. Edwardsburg, MI, USA

Janet A. Gilbertson, CSci FIBMS
National Amyloidosis Centre Royal Free and University College Medical School London, UK

Neil M. Hand, MPhil C.Sci FIBMS
Operational Manager Immunocytochemistry Histopathology Department Nottingham University Hospitals NHS Trust Nottingham, UK

J. Robin Highley, DPhil FRCPath
Clinical Fellow in Neuropathology Sheffield Institute for Translational Neuroscience Department of Neuroscience Sheffield University Medical School

Richard W. Horobin, BSc PhD
School of Life Sciences College of Medical, Veterinary and Life Sciences University of Glasgow Glasgow, UK

Toby Hunt, MSc BSc FIBMS
Laboratory and Mortuary Manager Department of Histopathology Great Ormond Street Hospital

Stuart Inglut, BSc (Hons)
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Peter Jackson, MPhil CSi FIBMS
Formerly Department of Histopathology and Molecular Pathology Leeds Teaching Hospitals NHS Trust Leeds Leeds, UK

Wanda Grace Jones, Ht(ASCP)
Immunohistochemistry Specialist Department of Pathology Emory University Hospital Atlanta, GA, USA

Laura J. Keeling
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Christopher Layton, PhD
Specialist Section Lead in Specimen Dissection Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Danielle Maddocks, BSc (Hons)
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Ann Michelle Moon, MSc MIBMS
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Guy E. Orchard, PhD C.Sci MSc FIBMS
Laboratory Manager Histopathology Department St. John’s Institute of Dermatology St. Thomas’ Hospital London, UK

Sherin Jos Payyappilly, FRCPath
Department of Histopathology Birmingham Heartlands Hospital Birmingham, UK

Anthony Rhodes, BSc MSc PhD CSi FIBMS
Professor Centre for Research in Biosciences Faculty of Health and Life Sciences University of the West of England Bristol, UK

Paul Samuel, BSc DMLT MIBMS
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Tracy Sanderson, FIBMS
Immunohistology Lead Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Lena T. Spencer, MA HTL(ASCP)QIHC
Senior Histotechnologist Norton Healthcare Louisville, KY, USA

Diane L. Sterchi, MS HTL(ASCP)
Senior Research Associate Histomorphometry Lead Department of Pathology Covance Laboratories Inc. Greenfield, IN, USA

John W. Stirling, BSc (Hons), MLett, AFRCPA, MAIMS, FRMS
Head of Unit The Centre for Ultrastructural Pathology Surgical Pathology – SA Pathology Adelaide, Australia

Jennifer H. Stonard, BSc (Hons), LIBMS
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Nicky Sullivan, CSci FIBMS
Department of Neuropathology and Ocular Pathology John Radcliffe Hospital Oxford, UK

S. Kim Suvarna, MBBS BSc FRCP FRCPath
Consultant Pathologist Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

Graeme Wild
Immunology Department Sheffield Teaching Hospitals Sheffield, UK

Anthony E. Woods, BA BSc (Hons) PhD MAIMS FFSc(RCPA)
School of Pharmacy and Medical Sciences University of South Australia Adelaide, Australia

Gregory Zardin, BSc (Hons) MSc MIBMS
Histopathology Department Sheffield Teaching Hospitals Sheffield, UK

General acknowledgments
Many Laboratory Scientists and Pathologists have contributed in different ways to the seven editions of this text and to acknowledge their individual advice and assistance is impossible. We express our thanks to everyone who has contributed since 1977. We owe Harry Cook special thanks for his advice and contributions to the earlier editions. Our thanks are also due to the colleagues we worked with in Nottingham and Sheffield during the production of this book.
We would like to thank all of our current authors, and those contributors whose previous work remains in some of the chapters in this new edition. Special thanks go to Richard Horobin who has contributed to all of the editions and to Marilyn Gamble for her work on the previous edition. Our thanks go to those who assisted in the preparation of the manuscripts and the production of the illustrations. We are grateful to Carol Bancroft for her considerable help with the editing and proof-reading.
Finally, we wish to thank the staff of our publishers for their unfailing help and courtesy.
John D. Bancroft, Kim Suvarna and Christopher Layton
Nottingham and Sheffield, UK

Acknowledgment to Alan Stevens
I have known Alan since he joined the Pathology Department at the University of Nottingham some 30 years ago. We had many discussions in those early years over whether the time had arrived for a multi-authored text on histological technique. It was apparent at that time that the subject was becoming too diverse for any single or two authors to cover in the depth that was required in the laboratories or the colleges where histotechnologists received their academic education.
In 1977 the first edition of this text was published and was due in no small part to Alan’s vision and diligent work in editing and even rewriting some of the chapters. His contributions to the succeeding editions were just as important and his medical knowledge was a significant factor in the development of the book. It has been a great pleasure working with him and I have greatly missed his contribution to the editing of this new edition, although much of his writing in the various chapters remains. The success over the years of Bancroft and Stevens owes a great deal to Alan Stevens. I wish to thank him and wish him well in his current and future medical education publications.
John D. Bancroft
Nottingham, UK
1 Managing the laboratory

Louise Dunk

Management is an integral aspect of the day-to-day life of the histopathology laboratory and is a major requirement of the accreditation process required by legislation in some countries. The accreditation standards include management as part of the evaluation and it is necessary that the laboratory worker is familiar with all of the processes involved. There are excellent books available which cover management issues in depth, and it is not the objective of this chapter to be a comprehensive guide to the subject. Rather, it discusses and concentrates on specific areas which have an impact on the operation of the laboratory; namely:

• Governance
• Risk management
• Quality management and establishing a quality system
• Personnel management
Other areas would include the management of estate, assets, equipment and supplies, business and budget management, and management of the scientific aspect and test repertoire of the service.
A pathology service may include a histology laboratory, an autopsy service, general cytology and a cervical screening/testing service. These areas will have many common management requirements but there will be some areas such as risk management where the issues will be individual to that section.
The surgical biopsy is sent for histopathological assessment to corroborate or dispute a clinical diagnosis by providing confirmation of data provided from other diagnostic tests. It should provide the clinician with valuable information on how to proceed with the treatment of the disease. Some resection specimens are taken as part of the treatment process, being referred to confirm the diagnosis, ensure adequate resection margins, to determine the extent of lymphatic/vascular involvement, staging, likely outcome and prognosis. Aside from simply determining a cause of death, autopsies may provide definitive data for a medical audit. They may be used to determine where medical procedures have been ineffective, or may give additional data for the future treatment of other patients. They can also provide a much needed final diagnosis and resolution for relatives.
Cytology samples are used as a screening process (e.g. cervical smears) and may assist in the early diagnosis of disease prior to the development of symptoms and thereby enable effective treatment. Cytology tests can also be used to monitor the stage of disease before/after treatment. This is accomplished by using non-invasive or minimally invasive techniques, which have a low risk of complications to the patient.


Risk management
Risk management is an essential and central part of all laboratory work. Organizations such as the Health and Safety Executive (HSE) and the Health Protection Agency (HPA) exist to ensure the safety of employees, patients and the general public in the United Kingdom. In the USA the Occupational Safety and Health Administration’s mission is to prevent work-related injuries, illnesses, and occupational fatality by issuing and enforcing standards for workplace safety and health, and most countries will have equivalent bodies and standards.
Regulations made under the Health and Safety at Work Act 1974 apply to all work situations, for example the Control of Substances Hazardous to Health (COSHH) Regulations and the Workplace (Health, Safety and Welfare) Regulations. The HSE enforces this act along with others, including the Health and Safety Offences Act 2008 . The overall message is that that employees are entitled to work in environments where risks to their health and safety are properly controlled (i.e. minimized). Under health and safety law, the primary responsibility is owed by employers, with employees expected to ensure their own safety, and that of their colleagues and/or patient’s by adhering to policies and procedures.
To comply with legislation and maintain accreditation, a laboratory must have an effective risk management policy. Any chance of something going wrong should be either negated or minimized, and therefore a laboratory’s risk management process should have procedures in place for:

• Identifying all risks that exist within the environment
• Assessing those risks for likelihood and severity
• Eliminating those risks that can be removed
• Reducing the effect of risks that cannot be eliminated
The pathology laboratory should have close links with, and feed into, the host organization’s risk management process. In most hospital laboratories, the laboratory manager will be accountable for risk management and the health and safety of the staff in their department, and often will be supported by a Risk Lead who will be responsible for the operational aspects of the system.
To function effectively and safely, all of a laboratory’s procedures and activities must be subjected to the risk management process. The risks in the laboratory are similar worldwide, albeit with a variation due to local circumstances. Health and safety and quality assurance incorporate a major aspect of risk management. All aspects of our working life incorporate a degree of risk and the risk management process allows us to prioritize, evaluate, and handle the risk appropriately. It is not possible to avoid or eliminate all risks, and in reality this may not be practical. It is important to identify and understand the risks that are involved in a laboratory’s working practices. An individual’s responsibility for risk management is dependent upon that individual’s role within the organization. The Chief Executive, for example, will be concerned mainly with risks associated with strategic issues affecting the organization as a whole and would only include histopathology within the risk assessment if it had a direct impact on these issues. Matters concerning the day-to-day running of the laboratory would not be of direct interest unless, of course, there was a significant reason for involvement such as major clinical or financial concerns or unmanaged risks or incidents, especially those likely to cause harm to patients, cause the organization to fail to achieve agreed targets or might attract adverse media publicity. A laboratory manager would be concerned with all risks associated with the department that they manage, but also how these might impact on other areas of the organization such as porters transporting samples or chemicals to the laboratory. They would also be required to alert the organization to the presence of risks which cannot be adequately controlled within or by the department.
The laboratory management team will deal with any laboratory-associated risk by ensuring that adequate resources are available to deliver the service, and by guaranteeing that the laboratory provides a service that is safe both for staff and patients. Staffing levels and competence, timeliness and quality of results, budgetary management, consumable and equipment supplies, and maintenance are some of the areas of concern. The laboratory management team must also ensure risk management procedures are in place for every aspect of a laboratory’s processes and environment.
The laboratory manager must ensure that day-to-day errors do not arise as a result of inadequacies in laboratory procedures and that quality control checks are in place to minimize the possibility of human errors: for example, a transcription error or mislabeling. Standard operating procedures (SOPs) should include COSHH data, risk assessments or equivalent, and also to include other health and safety information relevant to the procedure. This should include national legislation and guidance where available.
Scientific and support staff at the bench may be exposed to risks involving equipment malfunction due to poor maintenance or design. Poor-quality reagents may produce poor processing of tissues or inaccurate staining results. One of the most common accidents in the histopathology laboratory is the injury to fingers or hands from microtome blades or laboratory knives. It is the responsibility of each laboratory worker to reduce the risks associated with their day-to-day work by working in accordance with SOPs and associated risk and COSHH assessments. This will help ensure that everyone is working to the same standard and understands what is required to minimize risk to themselves, their colleagues and/or patients. It is importance that where risks are identified, the risk management measures which are put in place are regularly audited to assess whether they are being followed and are still appropriate and effective.

Risk identification
The risks within each laboratory section are best identified by the section lead and members of that team, working in conjunction with the laboratory’s health and safety lead. This ensures that the broadest possible spectrum of viewpoints is considered. During this process it is also useful to divide the risks into different categories, such as clinical, physical, chemical, infectious, etc., and even organizational, financial and political, depending on the area being risk assessed. For example a support worker unpacking the samples delivered to the laboratory might have noticed that more samples than usual have leaked. This could put both themselves and the porter at risk from infection and exposure to fixative, and if any of the contents has leaked beyond the specimen bag there could be a risk to other health workers and patients/visitors using the same route. This could just be a problem with one batch of specimen pots, but could also be a training issue for staff putting the samples in the pots. In raising the issue with their supervisor and giving them the opportunity to investigate the root cause, the support worker may have prevented harm to others and potential damage to the sample.

Risk analysis/evaluation
Analysis and evaluation of potential risks is an essential part of the process, and one that is used to identify both the likelihood and severity of these risks. By scoring the risks for likelihood and severity, it is then possible to use a matrix such as the one described below as a tool that will put a value on specific risks. This will then help prioritize them for further action.
The risk manager should put a system in place whereby all incidents and accidents are reported no matter how small. It is only by recording data that the full picture can be obtained and analysed and areas possibly overlooked initially be risk assessed and managed.

Severity and likelihood values
The following is an example of a severity scoring scale for incidents:

1. Low
• Minor injury or harm
• Minor loss of non-critical service
• Minor non-compliance with standards
• Minor out-of-court settlement
• Publicity mostly contained within organization. Local press coverage of no more than one day
2. Slight
• Injury or harm requiring less than 3 days absence from work or less than 2 days hospital stay
• Loss of service for less than 2 hours in a number of non-critical areas or less than 6 hours in one area
• Single failure to meet internal standards
• Civil action with or without defense, improvement notice
• Regulatory concern
• Local media coverage of less than 7 days
3. Moderate
• Medical treatment required and more than 3 days’ absence from work or more than 2 days’ extended hospital stay
• Loss of services in any critical area
• Repeated failures to meet internal standards or follow protocols
• Class action, criminal prosecution or prohibition notice served
4. Severe
• Fatality, permanent disability or multiple injuries
• Extended loss of essential service in more than one critical area
• Failure to meet national standards
• Executive officer fined or imprisoned, criminal prosecution – no defense
• Political concern, questions in parliament, national media coverage greater than 3 days
5. Catastrophic
• Multiple fatalities
• Loss of multiple essential services in critical areas
• Failure to meet professional standards
• Imprisonment of executive from organization
• Full public enquiry
Incidents may also be scored 1–5 for likelihood:

1. Incident unlikely to occur.
2. Incident likely to occur once in a 5-year period.
3. Incident likely to occur yearly.
4. Incident likely to occur once in a 6-month period.
5. Incident likely to occur once every 4 weeks or more frequently.
The risk factor is the severity multiplied by the likelihood of occurrence:

Very Low Risk – The majority of control measures in place or harm/severity small. Action may be long term.
Low Risk – Moderate probability of major harm or high probability of minor harm if control measures are not implemented. Action in the medium term.
Moderate Risk – Urgent action to remove or reduce the risk.
High Risk – Immediate action to remove/reduce the risk.

Risk management
The objective of the whole risk management process is to either remove or avoid risks, or manage them where removal is not an option. Removal would be possible, for example, by looking for alternatives to high-risk, harmful chemicals used in the laboratory. For example, prior to the 1970s, it was common practice to use mercuric chloride as a constituent of fixatives and, although this gave excellent quality fixation, it was extremely harmful to the environment and also to laboratory staff. Its use was subsequently stopped and alternative fixatives replaced it. Where risks remain, efforts should be made to reduce the effect or the possibility of the risk happening. The ways of controlling risk are numerous, but frequently there will be expert guidance or regulations issued by professional bodies or government agencies that the Risk Lead should ensure are implemented. Informal networking with professionals in similar laboratories can also provide valuable information and ideas as to how others have overcome the challenges of managing certain risks.
Audit is an essential tool in risk management. Regular audits of the effectiveness of the risk management measures put in place and the frequency and nature of incidents will allow the laboratory’s risk management team to assess them and amend and improve if required. Audit will also identify areas or tasks that may need more regular monitoring and may highlight training gaps for individuals or groups of staff. In addition, regular and targeted audits will provide evidence to assist with driving change should the risk be due to lack of funding for certain tasks or process or to processes outside the control of the laboratory management (e.g. labeling of samples in the operating theater).

Risk funding
Risk management should also consider insurance (individual or laboratory), although this is an important option. All medical staff carry medical liability insurance, which covers them in the event of any negligence claims. Similarly, professional indemnity insurance is commonly available today for non-medical laboratory staff who are much more at risk in today’s litigation-conscious society. The decision regarding whether or not to insure should be based on the risk assessment and the severity and likelihood of the risk. Some risks will not be appropriate for insurance cover for whatever reason, and in these instances the risk must be accepted by the organization.

Quality management
A quality management system is essential in order to provide the best possible service for the patient and clinicians. Quality is defined as a measure of how well a product or service does the job for which it is designed (i.e. conformity to specification).
Internal quality control of work processes is an important part of quality management, and has been the traditional way that bench work has been checked for many years. External quality assurance (EQA) schemes provide benchmarking against other laboratories and often provide access to best practice methods and expert advice on improving techniques/specific tests. However, a full quality management system should also encompass systems to ensure consistency, quality of service, confidence, standardization and continual improvement of all laboratory processes.
Quality management of a laboratory should ensure there are systems in place to monitor and improve areas such as organization and quality management systems. This will involve liaison with users, human resources, premises, the local environment, equipment management, information systems and materials. It will address the pre-examination process, the examination process, the post-examination phase as well as evaluation and quality assurance. Regular audit of the various components of the system will provide evidence of compliance with standards for accreditation. It should identify any trends and issues for concern, and confirm quality systems are working. Overall, all these measures should identify areas for quality improvement and show whether any improvements are working.

Accreditation is an important and long-established part of quality management in pathology laboratories. Accreditation allows confirmation that a department meets specific requirements for the users and clients, and fulfills appropriate legal requirements. The process will normally reduce risks from areas such as product failure, health risks, and company reputation. Many countries have their own long-established accreditation bodies and systems, but the International Organization for Standardization or ISO standards are being adopted by many countries as the standards they wish to work to and be accredited by. ISO is the world’s largest developer and publisher of international standards. There are ISO standards that cover many areas of activity, with the ones that affect medical laboratories being:

ISO 15189 – Medical laboratories – Particular requirements for quality and competence. This is the main standard that affects medical laboratories and that the majority will seek to become accredited to.
ISO 17043 – Conformity assessment – General requirements for proficiency testing. This standard specifies general requirements for the competence of providers of proficiency testing schemes, which would include external quality assurance schemes.
ISO 17011 – Conformity assessment – General requirements for accreditation bodies accrediting conformity assessment bodies . In order to assess and accredit laboratories according to ISO standards within their own country, national accreditation bodies such as CPA in the UK must themselves be accredited under this standard.
Laboratories wishing to be accredited must demonstrate a robust quality management system and consistent application of the standards, usually by undergoing assessment and surveillance visits from the accrediting body, plus providing annual reviews of the quality management system.

Quality control (QC)
This system checks that the work process is functioning properly. It includes processes utilized in the laboratory to recognize and eliminate errors. It ensures that the quality of work produced by the laboratory conforms to specified requirements prior to its release for diagnosis. Errors and/or deviations from expected results must be documented and include the corrective action taken, if required. In the laboratory, quality control has long been a component of accreditation requirements and should be ingrained in scientists as a daily practice.
Most laboratories have experienced scientists and support staff who have the responsibility of performing routine quality control checks prior to the release of slides for diagnosis. This QC evaluation will include, but is not limited to: accurate patient identification, fixation, adequate processing, appropriate embedding techniques, acceptable microtomy, unacceptable artifacts, and inspection of controls to determine quality and specificity of special staining and immunohistochemistry methods. Criteria should be established that would trigger a repeat if the QC findings were qualitatively or quantitatively unacceptable. Despite having a conscientious QC system in the laboratory, pathologists (having a higher level of expertise) perform the final QC examination as they assess/report the slide. It is their responsibility to determine that this is adequate for diagnostic interpretation. However, all personnel are responsible, such that errors and incidents should be recorded and audited regularly to identify trends. This will highlight any training needs and gaps.

External quality assurance (EQA)
In addition to local data collection and monitoring for internal quality control, external mechanisms provide valuable information regarding quality and peer comparisons and as an educational tool. In the UK, quality assurance of laboratory techniques is organized on a national basis. It is a system of peer review and registration with appropriate (approved) schemes. The non-profit-making NEQAS (National External Quality Assurance Scheme) organizes programs for histochemistry and immunohistochemistry.
In the USA, the National Society of Histotechnology (NSH) in partnership with the College of American Pathologists (CAP) (2006) , created the Histology Quality Improvement Program (HistoQIP). Their system scores each slide, assessing the fixation, processing, embedding, microtomy, staining and coverslipping. Additionally, CAP establishes national surveys for immunohistochemistry.
The UK quality assurance schemes were started by members of the profession to establish quality standards within histopathology. Registration with the schemes is now a requirement for accreditation. The quality assurance process is based on peer review of the stained sections submitted by participating laboratories. There are also medical quality assurance schemes for pathologists that cover many of the sub-specialties of histopathology.
The quality assurance schemes currently used in the UK are coordinated under the auspices of UK NEQAS and within this organization there are two individual schemes for histopathology, the NEQAS for immunohistochemistry and the NEQAS for cellular pathology techniques. The immunohistochemistry scheme gives participants the option to be assessed on general antibody panels, or more specialist laboratories may choose to participate only in the lymphoma or breast specialist areas. The cellular pathology scheme is subdivided into general, veterinary and neuropathology.
Accreditation standards require action be taken by poor performers to improve the quality of their preparations. Most schemes offer expert assistance and advice to laboratories that fall below the defined acceptable score.

Organization and liaison with users
An appropriate management structure for the department should exist so that the main functions can be adequately delivered. Staff at all levels should be qualified and trained for the work that they do and hold appropriate registration, if required. Competencies for the tasks performed should be regularly assessed, checked and recorded.
Many departments publish a mission statement outlining their business and aims. The quality objectives need to be documented so that all have clear objectives outlining who is responsible for achieving them and when they should be achieved by.
A laboratory will have many users, including patients, clinicians and those purchasing its services. It is essential when planning and developing a laboratory service that all users are consulted. In short, the department must know about the service it is providing/will provide. Likewise when monitoring the effectiveness and quality of a service, user feedback should be sought so that the service can be properly evaluated. Any complaints/praise should be followed up immediately, and should feed into the quality management system.

Premises, equipment and materials
The laboratory environment and equipment must be fit for all laboratory processes. Managers should ensure that there are adequate basic facilities for staff to do their jobs, such as rest and toilet facilities, adequate lighting, IT provision and space. There should also be enough space for equipment and storage. Equipment should be functional and be regularly maintained for safe use.
Staff must be trained and competent (in their own areas) to use all of the equipment and materials in a safe and effective way. Materials and equipment must be managed with regard to stock control and servicing. Procurement policies should ensure that quality stock is purchased, being fit for purpose and value for money.

Examination procedures
Any laboratory’s testing procedures may be multiple and complex, and in many laboratories its staff are required to rotate between sections. Also, it is essential that the methodology for all procedures and tests are documented in standard operating procedures (SOPs) to permit all staff to operate in a standardized and appropriate way. SOPs should cover all aspects of the testing process, from delivery of samples or reagents to the issuing of the final laboratory report. The SOPs therefore include not only the laboratory procedures but also those carried out by pathologist and clerical staff. It is important that SOPs that impact on areas or staff outside of the laboratory (e.g. porters delivering samples from operating theaters) are shared with the other departments responsible for managing that part of the process.
Accreditation standards require that SOPs and other policies are controlled within a document control system. This is usually a central database that holds authorized copies of documents, with controls on who can modify the data. The document control system must also ensure that only authorized and up-to-date copies of SOPs and policies are being used by staff performing the tasks. Any changes to a procedure must be captured within a further updated SOP. This must then be issued and any old SOPs removed from circulation.

Continuous quality improvement (CQI)
This is the system that is used proactively, to approach and identify opportunities to improve quality, before problems occur. It operates through evaluation and audit of all systems and processes in the laboratory. The goal is to improve care and safety for patients and staff through recognition of potential problems and errors – before they can occur. Good managers now realize that often failures, errors, and problems are usually due to the system processes and not necessarily the fault of the employee(s).
Regular and thorough audit of the many components of the laboratory’s quality management system and performance should be mapped against accreditation standards. This will help highlight any problem areas. Feedback from users provides useful information when evaluating the effectiveness and quality of the service. Any criticism received may well prompt an unscheduled audit of that particular part of the system.
CQI should include auditing of the laboratory’s procedures against not only accreditation standards but also those of the host organization/other services. Any audit findings that show that the laboratory’s processes are not adequate should result in corrective actions. These audit findings might also highlight improvements for processes, documentation, staff training or monitoring aspects of competency. Any corrective actions required should be completed as soon as possible in order that the service required by the users can be improved and brought up to standard quickly. CQI is a continuous cycle of audit and assessment of the service. If not monitored regularly, quality standards can slip as staff, equipment and reagents change. It is useful for the manager to establish an audit calendar to ensure that all areas are audited regularly with particular attention to ‘problem areas’.

Personnel management
One of the most important assets for a histology laboratory is its staff or personnel. More than any other pathology specialty, the laboratory process in histology is a very manual procedure, from sample receipt, through dissection (grossing), embedding, sectioning and staining. Many techniques are still reliant on skilled personnel rather than automation, and the laboratory manager must ensure that the department is staffed by an appropriate number of staff with the right level of skills to ensure that the process is robust, safe and cost-effective.

The role of the laboratory manager in staff management
The laboratory manager is accountable for the service provided by the laboratory, and should have the appropriate qualifications experience to undertake this task. As well as being the lead scientist for the department, laboratory managers are usually responsible for recruiting the appropriate staff, and also managing the human resource needs and professional direction of their staff. All staff should have comprehensive job descriptions so that they and their manager and supervisor know what is expected from them and to whom they are accountable. They should also have contracts that specify the terms and conditions that they are employed under.
Staff should have access to basic facilities such as handwashing, toilets and rest rooms. The manager should ensure that adequate breaks are allowed, especially in areas where staff cannot easily break off from what they are doing due to the high levels of concentration required, or because they work in areas where personal protective equipment is needed due to chemical or biological hazards. The European Working Time Directive and United States Department of Labor give guidance on how long staff can work without a break and maximum working hours per week.
The manager must ensure that there are appropriate numbers of staff with the required education, qualifications, training and competence to provide the service required. Managers must also ensure that staff have access to further education as required in order to continue to keep up with the latest knowledge and techniques related to the service being provided. The competency of staff to do the tasks within their job description needs to be assessed at regular intervals, and this together with regular formal appraisals should ensure staff are supported and provided with what they require to fulfill their roles. The manager must also address any issues with discipline or excessive absence from work to ensure that the workforce team functions optimally.
Regular staff meetings should be held that involve all levels of staff, in order that any new information can be passed on, such as new procedures or updates related to the risk and quality management systems. Regular meetings also allow staff to feed back any information they have or raise queries, and gives them access to supervisors or managers that they may not easily get during their routine day. Management techniques such as ‘Lean’ encourage short staff meetings at the start of each day so that any issues related to the days work can be raised and planned for, e.g. staff absence, workload, or other factors that might interrupt or disrupt the workflow.

Staffing the laboratory
Ensuring the right number and level of staff depends on the manager having a good understanding of the volume and complexity of work received. Good information systems are essential for recording and analyzing the work performed in a laboratory every year, and for understanding trends in workflow and complexity.
Guidelines such as those issued by the Royal College of Pathologists and the Institute of Biomedical Science in the UK advise what level of laboratory duties may be undertaken by which grade of staff, and have their own training and examination systems to enable consultant and postgraduate scientist staff to gain the qualifications they require. Scientific staff working in accredited laboratories in the UK should be registered by the Health Professions Council.
In the USA the National Accrediting Agency for Clinical Laboratory Sciences (NAACLS) fully accredits about 479 programs for medical and clinical laboratory technologists, medical and clinical laboratory technicians, and related professions. Other nationally recognized agencies that accredit specific areas for clinical laboratory workers include the Commission on Accreditation of Allied Health Education Programs and the Accrediting Bureau of Health Education Schools . Some States require laboratory personnel to be licensed or registered. Licensure of technologists often requires a bachelor’s degree and the passing of an exam, but requirements vary by State and specialty. Scientists may also gain certification by a recognized professional association, including the Board of Registry of the American Society for Clinical Pathology , the American Medical Technologists , the National Credentialing Agency for Laboratory Personnel , and the Board of Registry of the American Association of Bioanalysts .
Once the level and complexity of the workload is known, the workforce can be profiled to match its requirements, remembering that to be cost-effective, tasks not requiring registered or licensed scientists should be performed by support staff where possible.

With thanks to Sheffield Teaching Hospitals NHSFT for their kind permission to adapt and use the risk severity and likelihood values from the Trust risk policy.

Further reading

Accrediting Bureau of Health Education Schools (ABHES). website
American Medical Technologists (AMT). website
American Society for Clinical Pathology (ASCP) – Board of Registry. website
Board of Registry of the American Association of Bioanalysts (ABB). website
Clinical and Laboratory Standards Institute. Press release: From NCCLS to CLSI: One year. Online. Available at , 2006.
Clinical Pathology Accreditation (UK) Ltd Standards for the Medical Laboratory. Document name: PD-LAB-Standards v2.02 Nov 2010
College of American Pathologists (CAP). HistoQIP programme. Available at
Commission on Accreditation of Allied Health Education Programs (CAAHEP). website
DOH. Risk management in the NHS: D026/RISK/3M . London: Department of Health; 1994.
Health and Safety at Work etc Act. , 1974. Available at
Health and Safety Offences Act. , 2008.
Health Professions Council (HPC). website
Institute of Biomedical Science (IBMS). Managing staffing and workload in UK clinical diagnostic laboratories. Available at
ISO 15189. Medical laboratories – particular requirements for quality and competence . Geneva, Switzerland: International Organization for Standardization; 2007.
ISO 17011. Conformity assessment – General requirements for accreditation bodies accrediting conformity assessment bodies . Geneva, Switzerland: International Organization for Standardization; 2004.
ISO 17043. Conformity assessment –General requirements for proficiency testing . Geneva, Switzerland: International Organization for Standardization; 2010.
National Accrediting Agency for Clinical Laboratory Sciences (NAACLS). website
National Association of Histotechnology (NSH). website
National Credentialing Agency for Laboratory Personnel (NCA). website
Royal College of Pathologist (RCPath). Guidelines on staffing and workload in histopathology and cytopathology departments. second ed. Available at , 2005.
UK National External Quality Assessment Service (UKNEQAS). website
. Working with substances hazardous to health: what you need to know about COSHH, HSE leaflet INDG136(rev4). revised 06/09. Available at
Workplace (Health, Safety and Welfare) Regulations. HSE leaflet INDG244(rev2). Available at
2 Safety and ergonomics in the laboratory

John D. Bancroft
There are numerous workplace hazards in histology laboratories, and most countries have now passed regulations designed to improve this. These vary from country to country but the underlying theme is universal.
Risk management pertains not just to personal health and safety, but also to environmental health and safety. Hospital laboratories and research facilities have seen significant improvements in workplace conditions, but they remain contributors to environmental pollution.
The goal of this chapter is to lay out a risk management plan that is applicable worldwide. While general in scope to encompass a variety of regulations, it is specific regarding the hazards unique to histology. Most of the information is from Dapson and Dapson (2005) . Other references which should be in every laboratory include Montgomery (1995) , the Prudent Practices Series ( National Research Council 1989 , 1995 ), aids for preparing chemical hygiene plans ( Stricoff & Walters 1990 ), as well as guidelines from the Clinical and Laboratory Standards Institute concerning laboratory safety (2004) , biohazards (2005) and waste management (2002) . Indispensable publications from the Centers for Disease Control (USA) include guidelines for safety (1988) , HIV and tuberculosis 1990 , 1994) .

Risk management

Identify and evaluate hazards
The first step in risk management is to identify hazards in and emanating from the workplace. If this has never been done, it may be a formidable task, especially if there are old reagents or chemicals in poorly labeled containers. Anything that is unidentifiable or questionable should be set aside for disposal. Identification of hazards goes beyond making a chemical inventory, although that is a significant part of the effort. Electrical, mechanical and biological hazards are also included. In this initial identification stage, include the nature of the hazard(s) with the name, its location and the procedure(s) involved with its use. If no current use is found, then dispose of the item.
For hazardous chemicals, data sheets are available in most countries, and available from databases on the Internet. A file of data sheets should be kept in a secure location, and employees must be given reasonable access to it. It is also advisable to keep a duplicate file readily accessible in the laboratory in case of emergencies. Some reagents found in storage areas may be obsolete; sheets will be impossible to find for these. This creates a problem with no simple solution, because legitimate disposal may require having a data sheet, yet keeping the chemical also dictates that a sheet be on file. You will have to create one, or hire a qualified firm to do that for you.
Evaluate the severity of each of the hazards. What is the volume or magnitude of the hazardous item? How much is used per day (or some other meaningful unit of time)? Now put that information together with the data sheet. These are written for industrial-scale exposures, and you must weigh that against the scale of use in your laboratory. This evaluation must include risks associated with spillage and disposal as well as normal use. The hazards of a bulk container of formalin emptying onto the floor of a laboratory are quite different from spilling a 30 ml specimen container in a dermatologist’s office. Likewise, emptying hundreds of small formalin-filled specimen containers into a disposal drum or sink might present far greater exposure risk than handling each one during grossing. Do not underestimate risk, but keep the assessment proportional to scale and scope of operations.

Plan to minimize risk
Once the hazards have been listed and evaluated, decide how to reduce risk. Each item should be scrutinized, not just those offering the greatest dangers. Prioritize later. The goal is to reduce risks to acceptable levels, preferably through a cascading series of options that become progressively more burdensome and expensive. Work practice controls are the best way to tackle the problem; when pursued aggressively and with commitment at all levels of the institution, they usually are the only changes needed. Work practice controls involve eliminating, reducing and recycling everything possible. If they do not succeed, engineering controls should be implemented. These involve ventilation systems, fire protection devices and other expensive alterations to the facility. If all of these measures fail or are impossible to accomplish, personal protective equipment (PPE) must be used as a last resort. PPE should never be the first choice, although it may seem the most obvious way to protect workers.
There are several ways to reduce risk, the first of which should be to eliminate the hazard altogether. The list of obsolete chemicals in some of our labs is growing rapidly; how many does your laboratory still use? Remember using benzene and dioxane? No? Then do not be surprised in another few years to have histologists and biomedical scientists who never used xylene, toluene, chloroform, methacrylate, picric acid, uranyl nitrate and formaldehyde. A surprising number of laboratories are free of one or more of these highly dangerous substances and a few have eliminated all of them
Practically every hazardous chemical can be replaced today with a safer and technically superior substitute. The question is not whether it can be done, but if it might be done. The obstacle is rarely technical feasibility; most likely, it is human obstinacy. The notion that substitutes are not as good has been debased so often in everyday life that it is a wonder that it persists so strongly in the medical profession. Antifreeze, correction fluid, nail polish, hard surface cleaners, cosmetics, contact lens solutions and gasoline are just a few of the thousands of common materials in our lives that have undergone radical reformulation. In all cases, the products are safer, many work better, and some are less expensive.
If elimination of a hazard is out of the question, consider reduction. This will involve procedural changes, so be sure to weigh all implications before pushing ahead. A common idea for reduction is to use smaller specimen containers for fixation. Recycling is a final option for risk minimization. The volume in use at any given time might not be reduced, but the amount involved in storage and disposal will be cut drastically.
The plan must include justification to managers. Rationale for change should not rely solely upon improving safety. Financial considerations weigh heavily in any business, and could be your strongest argument for change. While many changes will cost more money initially, the long-term benefits are usually easy to calculate. For example, formalin substitutes are more expensive than formalin, but their use creates significant savings later. Workplaces and personnel do not need to be monitored for hazardous vapors, and disposal costs are usually reduced to zero. Less tangible but nonetheless real are the cost benefits of a healthier work force.

Implement the plan
Having a plan will do no good unless it is implemented. Prioritize the changes described in the plan. While easy changes should be tackled immediately, do not put off the challenging items that carry high health or environmental risk. Achieving financial gain quickly will help your cause, so be sure to include something at the outset with immediate positive economic impact.

Design standard operating procedures for working with hazards
Nearly all laboratories operate under a set of written, standard operating procedures (SOPs) mandated by a variety of accrediting or regulatory agencies. Detailed procedures for handling hazardous substances certainly should be central in these procedures, but other topics ought to be addressed as well. Personal hygiene practices should be a subconscious part of every workers’ behavior, but must be spelled out in the SOPs. Define the criteria for invoking the use of specific control measures, such as the use of protective equipment. Describe how to assure that fume hoods and other pieces of protective equipment are functioning properly. Make provisions for employee training, medical consultations and medical examinations. Detail spill procedures; define the kinds of spills that should be handled by laboratory workers and those too serious for anyone except trained HazMat responders. Establish a qualified officer or committee of qualified people to develop and administer these safety procedures.

Train personnel
Safety training is mandated by a variety of governmental regulations in several countries, and should be part of every department’s personnel practices. Trained people work more safely, efficiently and economically. In addition, the threat of employee litigation against the department is reduced. Regulations rarely address the issue of who should provide the training. In the past, it was common for one of the technical staff (usually the supervisor) to do this, that person obtaining the information as best he/she could. It is preferable, however, to have the trainer who is specially educated and experienced in health and safety matters.
Training must include general practices and may deal with very specific topics such as respirator use, handling select carcinogens, and working with formaldehyde. Each employee should sign a form verifying that training was received, a copy of which becomes part of the employee’s permanent record. The employee’s name, date and subject of training should be included on the certification form. Yearly retraining should be mandatory and documented. New employees, or employee’s assigned new hazardous tasks, must be adequately trained before beginning work.

Periodic reviews
On at least a yearly basis, all SOPs, risk assessments and training programs should be reviewed and updated as needed. Each written document should bear the date of creation and latest revision. Continue to minimize risks. Address any new risks that occur when different hazardous materials are brought into the workplace. Revise risk assessments and protocols to accommodate increased use of hazardous substances, especially as workloads increase.

Record keeping
Regulations often prescribe what records must be kept and for how long. It is prudent to record everything that pertains to regulatory compliance, risk assessment, causes and prevention of occupational illness or injury, employee health and safety training, exposure monitoring, occupational medical records, personal protective equipment and hazardous waste disposal practices. Records should be kept indefinitely, although 30 years past the duration of a worker’s employment is the term often prescribed by regulatory agencies. If in doubt, consider this: for how long would you want your estate to have access to health and safety records relating to your employment?

Occupational exposure limits
Most chemicals are hazardous to some degree; the question really is how hazardous are they? In other words, what would a safe level of exposure be? From many years of actual industrial experience, various agencies have developed standards for exposure to widely used chemicals. Generically, these are called occupational exposure limits, but each agency refers to its own values by unique names. OSHA’s Permissible Exposure Limits (PELs) are based upon scientifically based recommendations from the National Institute of Occupational Safety and Health, or NIOSH (2003) , but are also influenced by special interest groups and Congressional actions. OSHA limits therefore typically are more lenient. Another source of exposure limits, called Threshold Limit Values (TLVs ® ) is ACGIH ® , the American Conference of Governmental Industrial Hygienists (2004) . These limits are more widely used around the world for occupational standards.
An exposure limit is the maximum allowable airborne concentration of a chemical (vapor, fume or dust) to which a worker may be exposed. Presumably, it represents the concentration at or below which it is safe for most people to work; there will be individuals who react adversely below the limits because of hypersensitivity.
It is important to realize that exposure limits are properties of the worker and the workplace combined. They are not simply the maximum limits of vapor, fume or dust in the workplace; they are the maximum limits of exposure. This is especially important to consider when monitoring exposure levels. Monitor employees, not the workplace. Monitoring devices should be positioned as close as possible to the worker’s face in order to capture actual breathable quantities of hazardous material. For example, airborne levels of formaldehyde vapor a few inches above a grossing station’s cutting board may be much higher than concentrations at nose level, especially with well-designed ventilation.

Kinds of exposure limits based upon the duration of exposure

TWA (or TWAEV). The time-weighted average (time-weighted average exposure value) is the employee’s average exposure over 8 hours. Shorter exposures may exceed this value as long as the average exposure does not. There may be some short exposure that is too high for safety; that is covered below. When additional exposure is likely through the skin, that may be noted after the TWA. This is especially true for chemicals like phenol and methanol that pass quickly through skin.
STEL (or STEV). The short-term exposure limit (or value) is the highest permissible time-weighted average exposure for any 15-minute period during the work shift. It should be measured during the worst 15-minute period. The STEL is always higher than the TWA.
CL (or CEV). The Ceiling Limit (Ceiling Exposure Value) is the maximum permissible instantaneous exposure during any part of the work shift. Few chemicals are given both a STEL and a CL; the CL is usually reserved for highly dangerous substances.
For chemicals lacking either a STEL or CL, prudent values may be determined by multiplying the TWA by 3 for the STEL or by 5 for the CL, as is suggested by the Ontario (Canada) Ministry of Labor (1991) . When more than one harmful substance is present, complex formulas must be used to determine combined occupational exposure limits. These formulas are prescribed by various governments and vary from country to country.
IDLH. This airborne concentration is immediately dangerous to life and health. Chemicals with low IDLH should be considered very dangerous when spilled or when significant volumes are being dispensed. A single inhalation at or above this limit could have serious, if not lethal consequences.

Biological exposure indices
Can a worker determine if significant exposure has occurred? Can the chemical in question be detected in the worker by a clinical test? In a few instances, the answer is ‘yes’. ACGIH ® has established Biological Exposure Indices (BEIs ® ) as maximum values of analytes determined from clinical tests on exhaled air, urine or blood for a variety of hazardous chemicals, but only four are pertinent to histology: N, N-dimethylformamide, methanol, phenol and xylenes. Consult the latest booklet issued yearly by ACGIH ® for details on the first three chemicals.
Because xylene is used so pervasively in histology, and so many histologists are concerned with its effects, further information is presented here on this chemical. The isomers of xylene are metabolized to methylhippuric acids, which can be measured in exposed workers’ urine. The BEI ® for xylenes is 1.5 g methylhippuric acids per g creatinine. Samples are collected immediately at the end of a work shift.
BEIs ® are not intended to be used in diagnosing occupational illness. They are not maximum safe permissible values. Rather, they are to be used as indicators that workers may be exposed to significant concentrations of harmful substances, particularly if a worker or a group of co-workers repeatedly show values of the analyte at or above the BEI ® . For xylene in a well-ventilated histology lab, high methylhippuric acids in urine would probably indicate significant skin exposure.

Types of hazard
Systems of classifying the hazardous nature of chemicals range from simple pictographs with numerical ratings to comprehensive lists of formally defined terms. Even within a single country, government agencies may differ in how hazards are defined. While no single system will suffice worldwide, the following terms do have nearly universal meaning and should serve on a practical basis for describing the hazards encountered in histology. For convenience, hazards are first divided into two broad categories, health and physical. The latter certainly have ramifications for health, but present more immediate problems for storage, handling and building codes.

Biohazards can be infectious agents themselves or items (solutions, specimens or objects) contaminated with them. Anything that can cause disease in humans, regardless of its source, is considered biohazardous, even if the disease primarily occurs in animals. In many countries, biohazardous materials are specially labeled and disposal is generally strictly controlled.
Irritants are chemicals that cause reversible inflammatory effects at the site of contact with living tissue. Most often, eyes, skin and respiratory passages are affected. Nearly all chemicals can be irritating given sufficient exposure to tissue, so general hygiene practices dictate that direct contact be avoided as much as possible.
Corrosive chemicals present both physical and health hazards. When exposed to living tissue, destruction or irreversible alteration occurs. In contact with certain inanimate surfaces (generally metal), corrosives destroy the material. A chemical may be corrosive to tissue but not to steel, or vice versa; few are corrosive to both.
Sensitizers cause allergic reactions in a substantial proportion of exposed subjects. Nearly any chemical may cause an allergic reaction in hypersensitive individuals, so the key here is the prevalence of the reaction in the exposed population. True sensitizers are serious hazards, because sensitization lasts for life and only gets worse with subsequent exposure. It may occur at work because of the high exposure level, but chances are the chemicals will also be found outside the workplace in lower concentrations that aggravate the allergy. Formaldehyde is a prime example. Its vapors come off permanent press clothing, draperies, upholstery, wall coverings, plywood and many other building materials.
Carcinogens: While many substances induce tumors in experimental animals exposed to unrealistically high dosages, officially recognized carcinogens must present a special risk to humans. Criteria for the carcinogenic designation differ slightly among agencies, but in the end, any carcinogenic chemical used in histology is universally recognized as such. Examples include chloroform, chromic acid, dioxane, formaldehyde, nickel chloride, and potassium dichromate. Additionally, a number of dyes are carcinogens: auramine O (CI 41000), basic fuchsin (pararosaniline hydrochloride, CI 42500), ponceau 2R (ponceau de xylidine, CI 16150) and any dye derived from benzidine (including Congo red, CI 22120; diaminobenzidine and Chlorazol black E, CI 30235).
Toxic materials are capable of causing death by ingestion, skin contact or inhalation at certain specified concentrations. These concentrations vary slightly according to the agency making the designation, but differences are insignificant to the histologist. Some countries use the term poison when referring to this category. Toxic chemicals pose an immediate risk greater than the previously covered hazards, and some are so dangerous that they are given the designation highly toxic . Methanol is toxic; chromic acid, osmium tetroxide and uranyl nitrate are highly toxic. Use extreme caution when handling toxic substances; avoid highly toxic ones if possible.
Chemicals causing specific harm to select anatomical or physiological systems are said to have target organ effects . These are particularly dangerous substances because their effects are not immediately evident but are cumulative and frequently irreversible. There are numerous histological relevant examples: xylene and toluene are neurotoxins and benzene affects the blood. Reproductive toxins are especially prevalent (chloroform, methanol, methyl methacrylate, mercuric chloride, xylene and toluene, to name a few) and may warrant special consideration under occupational safety regulations of some countries.
The remaining hazard classes pertain to physical risks. Combustibles have flash points at or above a specified temperature. Flash point is the temperature at which vapors will ignite in the presence of an ignition source under carefully defined conditions using specified test equipment. It is a guide to the likelihood vapors might ignite under real workplace conditions. Flash point is not the temperature at which a substance will ignite spontaneously. Different countries and various agencies within those countries have their own unique values for the specified temperature. In the USA, OSHA defines it as 38°C, while the Department of Transportation uses 60.5 °C. Combustible liquids pose little risk of fire under routine laboratory conditions, but they will burn readily during a fire. It is better to choose a combustible product over a flammable one if all other considerations are equal. Clearing agents offer this choice.

Flammable materials have flash points below the specified temperature discussed above, and thus are of greater concern. Vapors should be controlled carefully to prevent buildup around electrical devices that spark. Special provisions for storage are usually mandated by national regulations, but local codes may impose even stricter measures. Storage rooms, cabinets and containers may have to be specially designed for flammable liquids; volumes stored therein may also be limited. Original manufacturers’ containers should be used whenever possible, and preferably should not exceed 1 gallon (4–5 liters).
Explosive chemicals are rare in histology, the primary example being picric acid. Certain silver solutions may become explosive upon aging; they should never be stored after use. In both cases, explosions may occur by shaking. Picric acid also forms dangerous salts with certain metals, which, unlike the parent compound, are potentially explosive even when wet. The best defense against explosive reagents is to avoid them altogether; this is certainly feasible today with picric acid.
Oxidizers initiate or promote combustion in other materials. Harmless by themselves, they may present a serious fire risk when in contact with suitable substances. Sodium iodate is a mild oxidizer that poses little risk under routine laboratory conditions. Mercuric oxide and chromic acid are oxidants that are more serious. Organic peroxides are particularly dangerous oxidizers sometimes used to polymerize plastic resins. Limit their volume on hand to extremely small quantities. Pyrophoric, unstable (reactive) and water-reactive substances are not generally found in histology. All involve fire or excessive heat.

Control of chemicals hazardous to health and the environment

Personal hygiene practices
There must be no eating, drinking or smoking in the lab. Application of cosmetics other than hand lotion likewise has no place within the laboratory setting. Wash hands frequently, but keep skin supple and hydrated with a good lotion. If hazardous powders have been handled, wash around your nose and mouth so that adherent particles are not ingested or inhaled. Solutions must never be pipetted by mouth.

Every chemical should be labeled with certain basic information; proper labeling of all containers of chemicals is mandated in some countries. Most reagents purchased recently will have most of the following already on the label, but older inventories may lack certain critical hazard warnings. Remember that solutions created in your laboratory must be fully labeled. Minimum information includes:

• chemical name and, if a mixture, names of all ingredients;
• manufacturer’s name and address if purchased commercially, or person making the reagent;
• date purchased or made;
• expiration date, if known;
• hazard warnings and safety precautions.
When putting a reagent’s name (or names of ingredients) on the label, use terminology that will be useful to those needing the information. In histology, we have many reagent names that are unfamiliar to chemically knowledgeable people who might be involved in an emergency. This is why it is so important to list ingredients, using names with widespread acceptance in the general field of chemistry; for example, use formaldehyde for formalin, acid fuchsin and picric acid for Van Gieson’s, and mercuric chloride, sodium acetate and formaldehyde for B-5.
Commercial products in their original containers will have the name and address of the manufacturer or supplier. If you put the material into another container, even ‘temporarily’, include this information on the new label. Chemicals in ‘temporary’ storage conditions have a bad habit of remaining there for years after laboratory personnel have moved on.
If the reagent is made in the laboratory, indicate who made it and when. Traceability could be critical if other information is lacking, as in the case of a Coplin jar of ‘silver stain’ left in a refrigerator. Is the solution one of those that is potentially explosive? Does anyone know which silver solution it is?
Many laboratories use small self-adhesive labels that say ‘Received: ——’. These are dated and affixed to each incoming container. Similarly, an expiration date should also be included for those chemicals that do not have an indefinite shelf life. Most inorganic compounds and many non-perishable organic chemicals are good for many years, but mixtures frequently deteriorate in a shorter time. Information on shelf life is hard to come by, and the best source is your own experience since each laboratory has different conditions and perhaps slightly varied formulations. Kiernan (1999) has included shelf-life data from his own extensive experience, which should serve as a good first approximation for your use.
Hazard warnings at a minimum should include the designations listed in the preceding section. This is the simplest and least ambiguous system. Pictographs (flames, corroding objects, etc.) are not universally recognized; some are obscure as to their meaning. Hazard diamonds are popular but carry risks of misinterpretation, especially since there are several systems in use. When there is an emergency, people may not think clearly or have time to figure something out. They need immediate access to the nature of the danger, and nothing provides that so effectively as the printed word. Briefly worded safety precautions may be appended to the hazard warning: for example, irritant, avoid contact with skin and eyes.
A multicultural workforce, not all of who may have the same native language, staff many labs. It is prudent to accommodate their needs by providing multilingual hazard warnings. Again, in an emergency you want no impediments to prompt and correct action.

Warning signs
Various countries have established different guidelines or mandatory regulations involving signage, so specific recommendations cannot be given here.

Protective equipment
There are certain general guidelines for clothing suitable for laboratory work that should be considered before protective equipment. Secure, close-toed footwear should be mandated; open-toed shoes and sandals offer no protection against spills or dropped items. While nearly all fabric today is resistant to destruction by histological solvents, such was not always the case. Certain early acrylic and acetate fibers dissolved almost instantly when in contact with xylene or toluene, creating a great deal of embarrassment when tiny drops of solvent hit the cloth. The possibility that such fabric still exists is real enough to take heed.
Aprons, goggles, gloves and respirators are the personal protective equipment (PPE) most likely to be used in the histology laboratory. In some countries, law for certain hazardous situations requires specified PPE. The following set of recommendations should be a routine part of general laboratory hygiene and will satisfy the most stringent regulations. When specific requirements exist for certain chemicals, such information will be included below in the section detailing common histological reagents. Aprons should be made of material impervious to the chemicals being used. Simple disposable plastic aprons are usually quite satisfactory, although heavy rubber aprons may be warranted when handling concentrated acids. Cloth laboratory coats are suitable only for protection against powders or very small quantities of hazardous liquids. Do not use them for protection against formaldehyde.
Goggles should be chosen specifically for each worker to accommodate the diversity of facial shapes and prescription glasses. Goggles not only come in a variety of sizes and shapes but also they are made for different functions. Choose only vented splashproof goggles for routine work in histology. These allow for ventilation, which reduces bothersome fogging of the lenses, but the vent holes are baffled so that splashing liquids are not likely to reach the eyes. Never cut holes in goggles to improve ventilation, as this defeats the protective function of the equipment. For severe conditions of exposure, wear a faceshield over splashproof goggles; never use a faceshield without the goggles. Finally, safety glasses are no substitute for goggles when handling hazardous liquids.
The issue of contact lenses arises frequently in discussions about eye protection (American College of Occupational and Environmental Medicine 2003). If liquids with no irritating fumes are being handled, contact lenses may be used safely in conjunction with appropriate goggles. Conventional goggles offer no protection against harmful vapors, which can become trapped beneath the lenses, causing corneal damage. If your eyes sting or water, your inhalation exposure is almost certainly beyond permissible or prudent limits. Regardless of whether you wear contact lenses, do not work under those conditions, even for brief periods.
Gloves are the most controversial PPE, and misinformation abounds. It is important to understand how gloves work, so that informed decisions can be made about glove selection. Glove material is rarely completely impermeable; it delays penetration of harmful material for a time sufficient to provide adequate protection. Chemical resistance refers to how well material holds up in the presence of solvents, but says nothing about how readily substances move through the material. In most cases, liquids rarely penetrate intact glove material. The vapors are the problem, both because they penetrate more efficiently through gloves and skin, and because the worker usually cannot detect them. Reputable manufacturers of gloves evaluate their products in standardized tests, measuring the time it takes for detectable amounts of a particular chemical to appear on the far side of the material. This is called the breakthrough time, and it increases non-linearly with glove thickness. A glove twice as thick as another made from the same material will not have a breakthrough time that is double that of the thinner glove. Schwope et al. (1987) present the most comprehensive listing of data on this subject.
Latex is one of the most permeable of all glove materials. Thick (8 mil) rubber gloves have a breakthrough time of 12 minutes with formaldehyde solutions. Latex surgical gloves are so thin (1.0–1.5 mil) that they offer no effective protection against formaldehyde or histological solvents. These gloves are suitable only for protection from biohazards. Keep in mind the startling increase in the incidence of latex sensitization, which has accompanied the widespread use of these gloves since the beginning of the AIDS epidemic.
Nitrile gloves are the best option for histological use. They are available in surgical-type thinness for brief intermittent exposures where fine dexterity is necessary. Exposures that are more serious can be safely tolerated with 8 mil nitrile gloves. Remember, however, that no glove material is effective against all classes of chemicals, and nitrile is no exception. Some chemicals in wide histological usage (xylene, toluene, chloroform) will permeate nitrile in seconds.
Respiratory protection against chemical vapors should rarely if ever be needed except in emergencies. Regulatory agencies stress that respirators are the protective equipment of last resort. No one in histology should be in a workplace whose vapor levels are even transiently higher than the PELs. Wearing respirators is uncomfortable, expensive and fraught with compliance hassles. Leave them to the people specially trained not only in respiratory use but also in dealing with such dangerous environments.
In the following discussion, the word ‘should’ is used, but substitute ‘must’ in countries having stringent respiratory protection standards. Workers should receive special training for wearing respirators because of the complexities of proper usage. Each worker needing this level of protection should be individually fitted with a respirator that exactly fits the contours of the face. The efficacy of the fit is then assured through a series of complicated tests that should be documented and repeated on a periodic basis. Workers should undergo medical evaluations and respiratory function tests to determine if they are physically qualified to wear respirators. Cartridges for respirators must be chosen carefully for the chemical environment. Both the type of chemical and the vapor concentration are vitally important considerations. Respiratory protection from airborne infectious materials is another matter altogether. Surgical masks are unacceptable because they fit poorly and have too large a pore size to filter out aerosols. HEPA (high efficiency particulate air) filters are suitable. Workers wearing HEPA masks may have to comply with applicable provisions of respiratory protection standards.

Ventilation is the foremost engineering control; ensuring proper airflow through a laboratory is the first critical step in improving working conditions. Every laboratory scientist should be aware of the following basic principles. For further details on hood design and placement, see Dapson and Dapson (2005) and Saunders (1993) . Laboratories should have two separate systems of ventilation, one for general air circulation (often combined with heating and air conditioning and called HVAC), and the other for local removal of hazardous fumes. They must work in concert to be effective, and must not merely shift the noxious vapors to another part of the facility.
General ventilation is for the physical comfort of the occupants of the room. Each hour, the entire volume of room air should be exchanged 4–12 times. That air should not contain significant quantities of hazardous vapors. If such vapors are originating somewhere in the room (from a grossing area, for instance), they should be dealt with at their source with an independent system of local ventilation.
Properly designed chemical fume hoods enclose the emission area, isolating it structurally and functionally from the rest of the room. A motor somewhere in the ductwork (preferably far from the hood) moves air directly to the outside. A sliding door (sash) usually fronts the system, and is an integral part of the way it works by controlling the face velocity of air entering the enclosure. It is a common misconception that high face velocities are good. In fact, strong airflow may create such turbulence inside the hood that contaminated air spills back out into the room. For vapor levels usually encountered in histology labs, a face velocity of 80–120 linear feet per minute is ideal. Control this by adjusting the height of the sash. As lifting the sash enlarges the opening, face velocity declines. Either a vaneometer built into the hood or obtained as an inexpensive handheld device measures face velocity. Always keep the sash at least partially open (unless the hood is designed to admit room air from another port) to prevent overtaxing the motor.
Improperly designed hoods will not be able to achieve optimal face velocity with the sash opened to a comfortable working height. Avoid these literally at all costs, for your facility’s money will only be wasted, giving you a false sense of security. There are important dimensional considerations that determine a hood’s effectiveness: the hood will develop dangerous eddies if too shallow and may not be able to move the full volume of air if too expansive.
There are other, external factors that influence how a hood works, and all center on the air supplied to the face of the hood. It should be obvious that a device removing air from a workplace must have a supply to draw upon. This is in addition to the amount required by the general ventilating system to exchange 4–12 room air changes per hour. Heating and air conditioning must also be balanced to account for the removal of air through the hood. Location of a hood is critical. Airflow into the face should be smooth and unimpeded. Surprisingly strong crosscurrents are generated by doors opening and closing, or by people walking by. Even the draft from general HVAC ducts can adversely affect hood performance. Any of these disturbances can draw harmful vapors out of the enclosure into the room, even against a net inward flow of air. Locate the hood, and by inference the hazardous work area, out of main traffic patterns and away from HVAC ducts.
Do not use fume hoods as storage or disposal devices. Objects within a hood disrupt airflow, and may block important air passages. Containers that emit vapors should not be placed within a hood except as a temporary safety measure. Remove the offending substance as soon as possible and put it into a secure container. Finally, do not put a waste chemical into a hood for evaporating it away unless there is no alternative in an emergency. Doing so is probably a violation of environmental regulations, and it may exceed the capacity of the hood to carry fumes away safely.
Ventilation devices other than fume hoods are used in histology labs; few are suitable unless vapor levels are already low. A non-enclosed system, such as a duct located above or behind the work area, may be powerful enough to draw contaminated air away from the worker as long as no crosscurrents are generated, but that is an unrealistic assumption. Workers must move about, and that usually destroys the effectiveness of unenclosed devices. Hoods that return air to the room after passing it through a filter may be suitable for localized workstations generating modest vapor emissions. Filters must be chosen with care. Vapor levels will dictate the size needed. Formaldehyde is not effectively captured by the filtration media used for solvent vapors. Filters become loaded and must be replaced, but how often this occurs is usually a mystery until odors are noticed out in the room. Since most workers in histology labs have impaired senses of smell, dangerous vapor levels may accumulate before anyone detects them. If filtration devices must be used, figure out how to determine effective life of the filters and establish a strict replacement regimen.
Air purification systems based upon ozone should not be used. They generate a chemical that is more hazardous than most of the fumes found in histology labs: ozone has a Ceiling Limit of 0.1 ppm according to ACGIH ® . Further, ozone from these purifiers does not seem to be effective in destroying formaldehyde vapors ( Esswein & Boeniger 1994 ).

First aid
With laboratory chemicals, the most common accidents requiring first aid are ingestion, eye contact and extensive skin contact. All health care professionals should have basic training in dealing with these situations at least and preferably with all aspects of first aid. Yearly safety training should include preparedness exercises on the most likely chemical accidents. Ingestion is encountered with patients and other non-laboratory staff, and frequently involves formalin. This is a tragic consequence of administrative neglect of basic safety issues. Laboratory chemicals should never be accessible to unattended patients, particularly those who because of age or illness are unable to think clearly about their actions. Improperly labeled containers are another cause of accidental ingestion. Patients should not be allowed to take fixed surgical specimens home; they present such a large risk from poisoning that no argument to the contrary is sufficient. If a body part needs to be specially cared for as part of a religious need, it should be given only to a responsible adult who can assure that it will never be accessible to unattended children.
First aid for ingestion of hazardous chemicals is not a simple matter. Some reagents will cause more damage if vomited and subsequently aspirated into respiratory passages; others are so toxic that the risk of aspiration is outweighed by the necessity to get the offending substance out of the body quickly. To solve this dilemma, some countries have established sophisticated networks of emergency response teams, which are admirably qualified to provide the best advice. If you have access to a Poison Control Center, or something similar, post the telephone number on each telephone in your laboratory. Time is of the essence in such emergencies, and preparedness may save a life. If enough people are available, get the victim to the emergency room while someone else contacts a Poison Control Center. If outside help is not possible, give a conscious victim a large quantity of water.
Splashing of dangerous chemicals into eyes is a common accident among those who fail to wear suitable goggles. Except for concentrated mineral acids, routine histological chemicals, including formaldehyde, are not likely to cause serious harm to eyes as long as proper treatment immediately follows an accident. All labs should be equipped with emergency eyewash stations, as either freestanding devices or small appliances affixed to sink faucets (the latter must be tested frequently to assure free flow of water). Current recommendations are to have such devices no more than 10 seconds or 30 meters from hazardous work areas. Ideally, the water temperature should be controlled to a range of 15–35°C. Portable eyewash bottles are not recommended and may be deemed unacceptable by regulatory agencies. These containers hold little liquid and may become contaminated with microorganisms.
Rinse the affected eye for 15–30 minutes, pulling the lids away from the eyeball. This is a seemingly interminable period, but do not shorten it. Emergency health care should be sought only after this treatment.
Treatment of skin contact with hazardous chemicals is simple: wash with water for 15–30 minutes. A quick rinse will not be sufficient for the more dangerous chemicals. Emergency showers should be as accessible as eyewash stations. If the substance is not readily water-soluble, use soap with the water wash. Immediately remove contaminated clothing, including wet shoes. Launder before wearing again, or discard the article. Formaldehyde-soaked leather will be difficult to salvage.

The advantages of using radioactive chemicals are rarely sufficient to justify their risks to health and the environment. Exceptions may pertain to therapeutic radioisotopes used as tracers. These emit very low levels of poorly penetrating radiation, and have half-lives measured in hours.
If radioactive substances are handled, a qualified radiation safety officer must oversee all aspects of the project, including waste disposal. Participating staff must be specially trained in radiation safety. This will allay fears as much as create a responsible work force. The work area should be monitored periodically with a radiation detector. Workers should wear dosimeters.

Storage of hazardous chemicals
Most laboratory chemicals can be safely stored in conventional cupboards. Dangerous liquids are best stored below countertop height to minimize the risk of bodily exposure in case a bottle is dropped and broken. Buy dangerous reagents in plastic or plastic-coated glass bottles whenever possible. Special storage provisions are warranted for acids, flammables, radioactive isotopes, controlled substances and hazardous chemicals in bulk containers.
Specialized acid cabinets are designed to contain the fumes emanating from most containers of strong mineral acids. They should be vented to the outside, using acid-resistant ductwork. Curiously, many of these storage devices contain some mild steel parts, which soon rust. Choose this equipment carefully for that reason. Paper labels on acid bottles should be checked periodically for corrosion that could lead to illegibility. Other special storage cabinets are usually mandated for all but the smallest quantities of flammable materials. These are designed to contain a fire within the cabinet. If they are vented, provisions must be made in the ductwork to prevent the spread of fire from the cabinet.
Certain flammable liquids present unusual fire and explosion risks because of their highly volatile nature and very low flash point. Isopentane and diethyl ether (‘ether’) are common examples. Opened containers cannot be resealed reliably. Never store these in a refrigerator or freezer unless these appliances are certified as suitable for an explosive atmosphere (mistakenly referred to as ‘explosion-proof’). The best advice is to avoid using these chemicals altogether. If that is not possible, buy only the quantity immediately needed, use it up if possible and do not try to store any leftovers.
Radioactive chemicals and controlled substances must be stored separately from other reagents. Cabinets should be locked. Access should be limited to a few specially qualified people.
Large containers present other risks. Even 5-gallon (20-liter) quantities can be too heavy for people to handle, especially for pouring operations. Equip these containers with spigots and keep the spigot above fluid level when not in use. Larger drums more than 200 liters require special handling equipment for moving and dispensing. Be sure any pumping device is completely compatible with the chemical. Avoid mild steel parts for fixatives and most plastics for xylene and toluene.
Transporting hazardous materials from storage to work areas can be risky. Carry glass containers with both hands, one hand beneath the jar or bottle. Rubber buckets should be used to carry highly dangerous materials like glass containers of mineral acids.

Spills and containment
Preparedness for spills begins with laboratory design. The goal is to prevent hazardous materials from reaching the outside environment. There should be no open floor drains unless they lead to special containment equipment that can be pumped out or drained by a hazardous waste hauler. Floor drains for showers can be built with a low dyke that prevents liquids on the floor from entering and keeps most of the water from the shower from getting out onto the floor.
How laboratory personnel respond to a spill will depend upon the nature of the hazard, the volume of the spill and the qualifications of the staff. Each chemical should be evaluated with these factors in mind. A gallon of alcohol spilled onto the floor presents a risk of fire but little health hazard, while the same quantity of formalin could be life threatening (20 ppm is imminently hazardous to life). Small spills are defined as those that can be safely handled by the immediate staff. Large spills present risks that surpass the qualifications of the same people to deal safely with the emergency and require specially trained HazMat or emergency response teams.
Develop plans for dealing with each family of hazardous material (acids, bases, flammables, etc.). Detail exactly what protective equipment is needed, and how each type of spill will be handled. Establish who will be called in the event the spill requires outside help, then contact them so they will be prepared. They may want an on-site visit to familiarize themselves with your facility’s layout, and certainly will want to discuss the types and magnitudes of hazards. Finally, train the laboratory staff on spill procedures and practice doing it with harmless material. Not having had the time to do this will prove to be a sorry excuse when the accident occurs.
If the amount of spilled material is limited to a few grams or milliliters, simply wipe it up with towel or sponge, protecting your hands with suitable gloves. Dispose of the towel or sponge appropriately; do not put it into the general trash, and protect the room from its vapors by sealing it within an impermeable plastic bag or other container.
In contrast to such very small incidents, an entirely different approach should be used for any other spills of dangerous materials. All personnel should evacuate the room or immediate vicinity of the problem. Assemble in a designated spot, and be certain everyone is there. On your way out, watch for your co-workers and assist anyone needing help getting out. Provide first aid if anyone has been splashed or is feeling the effects of vapors. Calmly discuss the magnitude of the spill and determine if it is large or small. There should be no mention of cause or blame here: it is immaterial to the immediate problem. If the spill is large, call an emergency response team and seal off the area. If small, decide how to handle the spill based on prearranged plans.
Spill neutralizing and containment kits should be available immediately outside the hazardous work area. These may be commercially purchased or assembled from common materials, and should include protective equipment and cleanup aids. Nitrile gloves similar in thickness to dishwashing gloves are adequate for most spills likely in histology; several sizes are available. Splashproof goggles and a faceshield are important. Provide disposable plastic aprons for chemical spills and disposable gowns for biohazards. If the staff is qualified and trained, equip the kit with respirators appropriate for the type of spill (a HEPA respirator for biohazards).
A good basic kit would also include cleanup items such as a dustpan and brush for powders, sponges, towels and mops for liquids, adsorbent material (vermiculite, kitty litter or a commercial sorbent), bleach (sodium hypochlorite) for biohazards, baking soda for acids, vinegar (5% acetic acid) for alkalis, and a commercial formalin neutralizing product. Have a sealable plastic bucket and heavy plastic bags for containment of the salvaged waste. Kits that are more sophisticated would contain instantaneous vapor monitoring devices so that the contaminated area can be checked before cleanup operations begin. Remember: the level of expertise of the staff will dictate how far to go with this.

One effective way to control hazardous chemicals is through recycling, as this reduces the quantities purchased, stored and discarded. Many clearing agents, alcohol and formalin can be recycled satisfactorily with proper equipment. Because these are the highest-volume chemicals in histology, the cost savings can be impressive, despite an initial outlay of capital funds.
Formalin is a mixture of volatile formaldehyde and nonvolatile salts in a solvent of water or water and alcohol (the small amount of stabilizing methanol can be ignored). Used solutions also contain solubilized and particulate components from the specimens. Through the process of simple distillation, water and formaldehyde are separated from all the other constituents. While that leaves the undesirable parts behind, the recycled product now lacks its salts and may not be at the proper concentration. Formaldehyde content can be assayed with a simple kit and adjusted as necessary. Fresh salts are readily restored to the solution.
Solvents should be fractionally distilled, as some of the contaminants in the waste are also volatile. Simple distillation will not separate these, and the resultant product may contain unacceptable amounts of water.
The most common problem with good distillation equipment is a foul amine odor detectable in the recycled product. It comes from deamination of protein in the waste during the distillation process. The freed amines evaporate readily and pass over with the other volatile components. Keeping the distillation chamber scrupulously clean is usually the key to avoiding the problem altogether. Formalin that has been heavily enriched with blood protein (as from fixing placentas) may require diluting with less bloody waste formalin. Pre-filter solutions contain many tissue fragments or coagulated protein.

Hazardous chemical waste disposal
Health care facilities are to prevent and treat illness, yet they are significant contributors to environmental harm especially in countries that have seen significant improvements in industrial pollution. On the other hand, if an effective pollution prevention program has been put into effect, there should be little waste to deal with. Reducing toxics use by substitution and minimization, coupled with recycling, could lead to amazingly low quantities of waste to be hauled away. The three highest-volume reagents, formalin, alcohol and clearant, can all be recycled. Formalin can be replaced with an effective glyoxal-based fixative that is drain-disposable in nearly all communities because of its ready biodegradability and low aquatic toxicity.
Options for disposal of hazardous chemical waste depend heavily upon national and local regulations, but the following recommendations should be valid anywhere. First, keep waste streams separated; do not mix different chemicals together unless told to do so by a qualified waste official. Second, know the hazard(s) of the waste. Is it flammable? Water-soluble? Toxic? Each of these factors affects the choice of disposal method. The best option for disposal is to pour the waste down the sanitary drain, from which it can be treated before entering the environment. Such waste, however, must not harm the biological processes that waste treatment facilities depend upon; nor must it pass through the system untreated. An example of the former would be formaldehyde in sufficient strength and quantity to kill off the bacteria driving the treatment process. Xylene typifies the latter, because its rate of biodegradation is too slow to be affected by the 1–3 day residence time in the wastewater treatment plant.
Formaldehyde is readily biodegradable. Nearly all organisms have an enzyme, formaldehyde dehydrogenase, which decomposes this chemical. The trick is to feed it into the system slowly enough so that it is diluted below toxic concentrations by the normal flow of water. Never dilute toxic material before pouring it down the drain, or follow disposal by ‘flushing with copious amounts of water’; this practice increases the volume passing through the treatment plant, which shortens the residence time and may impair biodegradation.
Work with your wastewater authorities, inform them of the nature of the waste (chemical composition and hazardous characteristics), and offer material safety data sheets. Propose a plan for disposal which includes the volume of waste, the time period over which it will be dumped, and the frequency of disposal. For example, you may wish to dispose of large quantities of waste formalin containing 3.7% or 37,000 ppm formaldehyde over a 1-hour period each working day. This could be accomplished by trickling the waste into a sink from a carboy equipped with bottom spigot adjusted so that it takes an hour to become empty. There would be no risk to the treatment plant at this flow rate of 2 ml/second. Dumping a large volume of waste all at once is not good because it tends to travel in a slug and may fail to become sufficiently diluted to protect the treatment plant.
Some waste can be rendered more acceptable for drain disposal. Acids and bases can be neutralized and formalin can be detoxified with commercial products. Be sure the pretreatment process is safe, effective and acceptable. Never attempt to detoxify formalin by mixing with bleach (sodium hypochlorite) or ammonia. Both reactions are exothermic and could quickly get out of control, spewing vapors and hot fluid all over the lab. Know what the reaction products are, and be certain that they are indeed suitably low in toxicity. Water-insoluble solvents are never drain disposable, even if purportedly biodegradable. If they are combustible with sufficiently high caloric value, they may be eligible to be mixed with the fuel in an oil-fired heating system in certain countries. Good candidates for this option are any of the clearing agents except the halogenated solvents chloroform, trichloroethane and their relatives. Note that this method is not burning the waste in an incinerator. The difference is subtle but important. An incinerator exists for the sole purpose of destroying waste, while a furnace provides heat.
If you cannot get rid of a waste by drain disposal or combustion in a furnace, you must resort to a waste hauler. In some countries, the waste generator (your facility) bears the ultimate responsibility and liability for the waste that someone else takes away and supposedly deals with properly. If you are generating waste that must be removed from the premises for burial or incineration, you should do everything possible to eliminate that chemical from your lab. There is no better advice; however, you want to view it, whether from a financial, health or environmental perspective.

Control of biological substances hazardous to health and the environment

An Exposure Control Plan for biohazards should be written. It may be part of the Chemical Hygiene Plan or an independent document, but definitely should be part of your SOPs. As with chemical hazards, workers should be trained initially, refreshed at least yearly and immediately introduced to any new procedures that present additional risks.

People who work in histology laboratories may not be exposed to quite the level of risk that many health care workers are, but they face hazards that may be more subtle. There are three potential routes of exposure: inhalation of aerosols, contact with non-intact skin and contact with mucous membranes (eyes, nose and mouth). Knowing how infectious agents can reach you is the foundation of protecting yourself and your co-workers. Practice universal precautions: handle every specimen as if it were infectious.
Fresh specimens of human origin must always be considered potentially infectious. Most animal tissue does not carry that risk, but there are important exceptions. Species known to be capable of transmitting disease to humans, and animals intentionally infected or known to be naturally infected with transmissible diseases, must be handled with the same precautions as would be used for human tissue.
The first and most obvious source of biological risk is with fresh tissue and body fluids; grossing carries the highest risk of all histological activities. Fixed specimens have a much-reduced risk because nearly all infectious agents are readily deactivated by fixation. Specimens must be thoroughly fixed for this to happen. Certain tissues like liver, spleen, placenta and lung do not fix well unless grossed thinly, and may remain raw (unfixed) in the center after days of exposure to the fixative. Those centers are potentially infectious. Some fixatives require more time than that available in rushed pathology labs, so tissues in the first several stations of a tissue processor may remain biohazardous. Complete penetration by alcohol will kill all infectious agents except prions, so it is safe that properly processed specimens are free from microbial risk and can be handled without special precautions.
Prions, the agents of spongiform encephalopathies like Creutzfeldt-Jakob disease, scrapie, chronic wasting disease and mad cow disease, present a more difficult challenge. Even normal steam sterilization fails to inactivate these particles, and common effective chemical treatments like sodium hypochlorite and phenol create artifacts in tissue. Histological specimens can be decontaminated by immersion in formalin for 48 hours, followed by treatment for 1 hour in concentrated formic acid and additional formalin fixation for another 48 hours. Rank (1999) reviewed the risks and decontamination protocols for histology labs.
Cryotomy presents special risks because tissue is usually fresh. Small dust-like particles generated from sectioning may become airborne, a risk vastly magnified with the use of cryogenic sprays. Do not clean the cabinet with a vacuum unless the device is equipped with a HEPA filter. Sterilize surfaces with chlorine bleach or a suitable commercial disinfectant; avoid formaldehyde solutions as these present a chemical risk to the person doing the cleaning. Good-quality latex or nitrile surgical gloves are perfectly acceptable protective devices for biohazards. Goggles should be worn during grossing anyway to protect against chemical splashes, and will do double duty against infectious agents. A faceshield may be warranted in some cases. Aprons or laboratory coats will keep clothes clean, but do not wear these used protective articles outside the laboratory (especially to the cafeteria!).

Disposal of biohazardous waste
Biohazardous waste should be incinerated on-site or hauled away. Either way, potentially infectious waste ought to be segregated from chemical and non-regulated waste, which may be barred from incinerators designed for biohazardous materials. Fixed wet specimens and their fluid are chemically hazardous and may be infectious. Together they pose a difficult problem.

Control of physical hazards

From equipment
Equipment may present risks from electrical and mechanical factors, which can be minimized by proper installation, care and personnel training. Keep a log for each piece of equipment, listing its installation date, the person and firm performing the installation, and the initial diagnostic test results that assure the item is working properly. Include a schedule for preventive maintenance and a complete service record is good laboratory practice.
Electrical shock most often arises from improperly grounded devices. Have a qualified person verify that all outlets are properly polarized and grounded. Plug equipment into outlets, not into extension cords.
Electrical equipment poses a risk of igniting flammable vapors. Nearly all switches may spark, including those associated with doors. Devices sold as ‘explosion-proof’ have their switches sealed to prevent contact with flammable vapors. Refrigerators and freezers must never be used to store highly flammable chemicals like ether and isopentane unless they are rated as suitable for flammable environments. Likewise, household microwave ovens should not be used to heat flammables because the door interlock switch may spark (this switch stops the magnetron when the door is opened). Some laboratory microwave appliances vent the chamber sufficiently to prevent potentially explosive vapor concentrations from building.
Today, mechanical dangers from histological equipment are generally confined to burns from hot surfaces. Most modern devices have adequate safety features that have eliminated some of the more common hazards encountered decades ago. If you use older equipment, be aware of its shortcomings. Many centrifuges have lockout devices that prevent the lid from being opened while the rotor is moving. Distillation equipment should be purchased only if it has safety features that include high temperature and low liquid volume cutoff switches. Specialized apparatus for electron microscopy should be used only by people specially trained in the inherent risks.
Devices that emit an open flame (such as Bunsen burners or alcohol lamps) must never be used in an environment where flammable solvents are present. Electrical appliances for heating or sterilizing are far safer and more convenient than a gas-fired or alcohol-fueled implement.
Broken glass and disposable microtome blades present risks, particularly if they are contaminated with chemical or biological material. Special ‘sharps’ containers are used for the disposal of such items. Microtomes and cryostats are particularly dangerous; be sure to remove blades before cleaning such equipment.

Hazards and handling of common histological chemicals
Extracting succinct information from data sheets and reference books is a time-consuming task that has served as a major impediment to designing proper training programs and labels. The following compilation includes most of the chemicals commonly used in histology laboratories on a routine basis. Permissible exposure values are from OSHA unless otherwise indicated as being taken from ACGIH ® (2004) or NIOSH (2003) . All IDLH values are from NIOSH and Biological Exposure Indices (BEIs) are from ACGIH ® . Many PELs have been revised downward since 2002. The listed hazards are applicable to the quantities normally handled on a laboratory scale and may be inappropriate for bulk quantities. Recommendations for glove material are from Schwope et al. (1987) . In reality, selection of glove material may have to balance chemical resistance against practicality for the tasks likely to be encountered. Some chemicals have been deemed essentially non-hazardous under normal laboratory conditions of use.
The intent of this section, and indeed this entire chapter, is to provide guidelines for a safe workplace. Most hazardous chemicals can be handled safely with a minimum of effort and equipment, but a few cannot. These have been identified clearly and should be eliminated or at least reduced to the smallest quantity possible. Suitable substitutes have been identified. The toxicology of most chemicals is not well known, and new information, usually damaging, continues to become available.

Acetic acid. TWA = 10 ppm; STEL = 15 ppm (ACGIH ® ); IDHL = 50 ppm. Irritating to respiratory system (target organ effects); concentrated solutions are severe skin and eye irritants, corrosive to most metals and combustible (flash point = 43°C). Avoid skin, eye and respiratory contact. Use a chemical fume hood, nitrile gloves, goggles and impermeable apron when dispensing concentrated acid. Do not use rubber (latex) gloves. Always add acid to water, never water to acid, to avoid severe splattering. Do not mix concentrated (glacial) acetic acid with chromic acid, nitric acid or sodium/potassium hydroxide. Dilute (1–10%) aqueous solutions are relatively benign.
Acetone. TWA = 1000 ppm (500 ppm ACGIH ® , 250 ppm NIOSH); STEL 750 ppm; BEI = 50 mg acetone/liter of urine at the end of the shift. Highly flammable (flash point = −16°C) and very volatile. Great risk of fire from heavy vapors traveling along counters or floors to a distant ignition source. Not a serious health hazard under most conditions of use but be aware that acetone can be narcotic in high concentration. Inhalation may cause dizziness, headache and irritation to respiratory passages. Skin contact can cause excessive drying and dermatitis. Moderately toxic by ingestion. Protect skin with Neoprene gloves.
Aliphatic hydrocarbon clearing agents. TWA = 196 ppm (manufacturer’s recommendation). Very low toxicity: non-irritating and non-sensitizing to normal human skin. Combustible (40°C) or flammable (flash point = 24°C). Limit skin exposure to minimize de-fatting effects. Neoprene or nitrile gloves are satisfactory. Recycle by fractional distillation, burn as a fuel supplement or use a licensed waste hauler for disposal.
Aluminum ammonium sulfate, aluminum potassium sulfate and aluminum sulfate. Not dangerous in laboratory quantities except as eye irritants.
Ammonium hydroxide. TWA = 50 ppm OSHA (25 ppm ACGIH ® ); STEL = 35 ppm as ammonia gas; IDLH = 300 ppm. Severe irritant to skin, eyes and respiratory tract. Target organ effects on respiratory system (fibrosis and edema). Wear rubber or nitrile gloves. Store away from acids. Do not mix with formaldehyde as this generates heat and toxic vapors. Spills of 500 ml or more may warrant evacuation of the room.
Aniline. TWA = 5 ppm (2 ppm ACGIH ® ) with additional exposure likely through the skin; IDLH = 100 ppm. A very dangerous reagent, which should not be used if possible. Moderate skin and severe eye irritant, sensitizer, toxic by skin absorption, carcinogen. Excessive exposure may cause drowsiness, headache, nausea and blue discoloration of extremities.
Celloidin (stabilized nitrocellulose). Harmless as a health hazard but dangerously flammable as a solid. May deteriorate into a crumbly, potentially explosive substance requiring professional assistance for removal. Solutions usually contain highly flammable ether and alcohol.
Chloroform. CL = 50 ppm; STEL = 2 ppm (NIOSH); IDLH = 500 ppm. Toxic by ingestion and inhalation. Overexposure to vapors can cause disorientation, unconsciousness and death. Target organ effects on liver, reproductive, fetal, and central nervous, blood and gastrointestinal systems. Carcinogenic. Practical glove materials are not available. This is one of the most dangerous and difficult chemicals in histology because workers in most laboratories simply cannot receive adequate protection from vapors and skin contact. Legitimate disposal may be very challenging. Do not burn. Do not evaporate solvent to the atmosphere. Avoid all use.
Chromic acid (chromium trioxide). TWA = 0.5 mg chromium/cubic meter (ACGIH ® ); CL = 0.1 (0.05 ACGIH ® ) mg chromium/cubic meter; IDLH = 15 mg chromium/cubic meter. Highly toxic with target organ effects on kidneys; corrosive to skin and mucous membranes; carcinogenic. Strong oxidizer. Avoid all skin contact. Nitrile, latex and Neoprene gloves are not suitable except for limited contact; suitable protective material not readily available or practical for laboratory use. Chromium is a serious environmental toxin. Drain disposal is not a legitimate option for any solution containing chromium, including subsequent processing fluids following fixation or rinses after staining procedures involving chromium. Give this chemical high priority for complete elimination from your lab.
Diaminobenzidine (DAB). Human carcinogen. Solutions pose little health risk under normal conditions of use. Disposal of DAB and subsequent rinse solutions down the drain creates environmental problems. These wastes can be detoxified with acidified potassium permanganate according to the methods of Lunn and Sansone (1990) ; see Dapson and Dapson (2005) for a simplified procedure. Do not use chlorine bleach, as the reaction products remain mutagenic ( Lunn & Sansone 1991 ).
Dimethylformamide (DMF). TWA = 10 ppm; additional exposure likely through skin contact; IDLH = 500 ppm. Eye, nose and skin irritant. May cause nausea. May be a reproductive toxin. Facilitates transport of other harmful materials through skin and mucous membranes. Combustible liquid (flash point = 136°F). Avoid all skin and respiratory contact. Use DMF only in a fume hood with suitable gloves (butyl rubber). Common glove materials do not provide adequate protection. Dispose of DMF only through a licensed waste hauler.
Dioxane (1,4-dioxane). TWA = 100 pm (20 ppm ACGIH ® ); additional exposure likely through skin contact; CL 1 ppm (NIOSH); IDLH = 500 ppm. Skin and eye irritant: overexposure may cause corneal damage. Readily absorbed through skin and mucous membranes. Delayed target organ effects in central nervous system, liver and kidneys. Only butyl or Teflon gloves are suitable. Flammable liquid, which develops explosive properties (peroxides) after a year. Do not recycle, as the risk of creating explosive peroxides increases greatly. Avoid all use of this chemical.
Dyes. There are thousands of dyes and many have been implicated in causing cancers in rats under highly unrealistic circumstances. All should be handled with due caution when in the powder state, but liquids pose little risks except through skin contact and ingestion. Dyes containing the benzidine nucleus are now considered known human carcinogens and must be treated accordingly in both handling and disposal. See the section on carcinogens earlier in this chapter for examples.
Ethanol. TWA = 1000 ppm. Skin and eye irritant. Toxic properties are not likely to be significant under intended conditions of use in a laboratory. Use butyl or nitrile gloves, not rubber or Neoprene. Flammable liquid. Recycle via distillation.
Ether (diethyl ether). TWA = 400 ppm. Mild to moderate skin and eye irritant. Overexposure to vapors can produce disorientation, unconsciousness or death. Target organ effects on nervous system following inhalation or skin absorption. Dangerously flammable liquid that forms explosive peroxides. It is extremely volatile and difficult to contain. Do not store in a refrigerator or freezer unless the appliance is rated for an explosive atmosphere. Use a licensed waste hauler. Because of the uncontrollable physical hazard, avoid use of this substance if possible.
Ethidium bromide. May be harmful by ingestion, inhalation or absorption through the skin. Irritating to skin, eyes, mucous membranes and upper respiratory tract. Chronic exposure may cause alteration of genetic material. Dispense powder under a fume hood and wear any type of gloves.
Ethylene glycol ethers (ethylene glycol monomethyl or monoethyl ether, Cellosolves). TWA = 200 ppm (5 ppm (ACGIH ® ); additional exposure likely through skin. Toxic by inhalation, skin contact and ingestion, with target organ effects involving reproductive, fetal, urinary and blood systems. Combustible liquids (flash point = 43–49°C). Avoid all use, substituting propylene-based glycol ethers. If substitution is not possible, wear butyl gloves and use a fume hood for all tasks involving these reagents.
Formaldehyde and paraformaldehyde. TWA = 0.75 ppm (0.016 ppm NIOSH); STEL = 2 ppm; CL = 0.3 ppm for 15 minutes ACGIH ® (0.1 ppm NIOSH); IDLH = 20 ppm. Severe eye and skin irritant. Sensitizer by skin and respiratory contact (this is the most serious hazard for most laboratory workers). Toxic by ingestion and inhalation. Target organ effects on respiratory system. Carcinogen. Corrosive to most metals. All workers exposed to formaldehyde should be monitored for exposure levels on a periodic basis. Exposure of the skin during grossing is the greatest risk in a well-ventilated lab. Latex surgical gloves are nearly worthless as protective devices. Thin nitrile gloves are better but cannot be used safely for extended periods. Recycle as much waste as possible by distillation and have the remainder taken away by a licensed waste hauler or detoxified by a commercial product. Drain disposal of limited quantities of formaldehyde may be permitted in some communities. Satisfactory substitutes are now available worldwide and offer substantial technical advantages.
Formic acid. TWA = 5 ppm; STEL = 10 ppm (ACGIH ® ); IDLH = 30 ppm. Mild skin and severe eye irritant. Corrosive to metal. Avoid skin, eye and respiratory contact; use a chemical fume hood. All common glove materials except latex are suitable. Always add acid to water, never water to acid, to avoid severe splattering.
Glutaraldehyde. CL = 0.2 ppm NIOSH (0.05 ppm ACGIH ® ). Severe skin and eye irritant; toxic by ingestion. Wear butyl or Neoprene gloves and use a hood.
Glycol methacrylate monomer. No established PELs. Sensitizer. Flammable liquid. Avoid all skin, eye and respiratory contact. Common glove materials are probably not suitable, based on information concerning other methacrylates. To avoid a dangerous exothermic reaction, do not polymerize large quantities of this monomer. Polymerize small quantities for disposal.
Glyoxal. No established PELs. Glyoxal solutions have no vapor pressure (do not give off fumes) and thus pose no inhalation risk. Irritant to skin and eyes. Ingestion may produce adverse fixative effects on the gastrointestinal tract. Wear nitrile gloves and goggles. Favorable ecotoxicity profile. An excellent substitute for formaldehyde-based fixatives.
Hydrochloric acid. CL = 5 ppm (2 ppm ACGIH ® ); IDLH = 50 ppm. Strong irritant to skin, eyes and respiratory system. Target organ effects via inhalation on respiratory, reproductive and fetal systems. Corrosive to most metals. Concentrated acid is particularly dangerous because it fumes. Use a fume hood, goggles, apron and gloves made of any common material except butyl rubber. Always add acid to water, never water to acid, to avoid severe splattering.
Hydrogen peroxide. TWA = 1 ppm; IDLH = 75 ppm. Solutions less than 5% are essentially harmless. Concentrated solutions are very hazardous and should not be used.
Hydroquinone. TWA = 2 mg/cubic meter; CL = 2 mg/cubic meter for 15 minutes (NIOSH). Irritant capable of causing dermatitis and corneal ulceration. Toxic by ingestion and inhalation. May cause dizziness, sense of suffocation, vomiting, headache, cyanosis, delirium and collapse. Urine may become green or brownish green. Lethal adult dose is 2 grams. All common glove materials are suitable except latex. Avoid contact with sodium hydroxide.
Iodine. CL = 0.1 ppm; IDLH = 2 ppm. Strong irritant and possibly corrosive to eyes, skin and respiratory system. Dermal sensitizer. Toxic by ingestion and inhalation. Wear nitrile gloves and use a hood when handling iodine crystals. Histological solutions are essentially harmless except if ingested.
Isopentane. TWA = 1000 ppm (600 ppm ACGIH ® , 120 ppm NIOSH); CL = 610 ppm for 15 minutes; IDLH = 1500 ppm. Excessive exposure to vapors causes irritation of respiratory tract, cough, mild depression and irregular heartbeat. Ingestion causes vomiting, swelling of abdomen, headache and depression. Chilled isopentane may freeze the skin but otherwise is harmless to it. Extremely flammable (flash point = −57°C) and highly volatile, making this a very dangerous chemical. Never store it in a refrigerator or freezer unless the appliance is rated for an explosive atmosphere. Protect hands from frostbite.
Isopropanol. TWA = 400 ppm (200 ppm ACGIH ® ); STEL = 400 ppm; IDLH = 2000 ppm. Mild skin and moderate eye irritant. Toxic by ingestion. Flammable liquid (flash point = 12°C). Practically harmless except for flammability under normal conditions of use. Recycle by fractional distillation.
Limonene. No PELs established. Generally regarded as safe as a food additive in minute quantities, but a dangerous sensitizer when handled as in histology. May cause respiratory distress if inhaled. Use a hood and gloves (butyl, Neoprene or nitrile). Clearing agents containing limonene usually cannot be recycled back to the original product because they also include non-volatile antioxidants and diluents.
Mercuric chloride. TWA = 0.01 mg mercury/cubic meter; additional exposure likely through skin contact; IDLH = 10 mg mercury/cubic meter. Severe skin and eye irritant; target organ effects on reproductive, urogenital, respiratory, gastrointestinal and fetal systems following ingestion and inhalation. Severe environmental hazard. Corrosive to metals. Avoid all use if possible because of the impossibility of preventing environmental contamination. Most processing solutions will become contaminated with mercury if any specimens have been fixed in B-5, Helly’s, Zenker’s or similar fixatives. Reagents used to ‘de-Zenkerize’ sections release mercury. None of these must be allowed to go down the drain. Legitimate disposal of mercury-containing waste is difficult and very expensive, if not impossible, in some areas of the world. Replace mercuric fixatives with zinc formalin or glyoxal solutions.
Mercuric oxide. Strong oxidizer. See mercuric chloride for other information.
Methanol. TWA = 200 ppm, STEL = 250 ppm (ACGIH ® ); additional exposure likely through the skin; IDLH = 6000 ppm. Moderate skin and eye irritant. Toxic by ingestion and inhalation, with target organ effects on reproductive, fetal, respiratory, gastrointestinal and nervous systems. May cause blindness or death. Flammable (flash point = 12°C) and rather volatile. Use butyl gloves; other common glove materials are ineffective. Recyclable.
Methenamine. No PELs established. Powder may cause irritation; solutions pose little risk under normal conditions of use.
Methyl methacrylate monomer. TWA = 100 ppm (50 ppm ACGIH ® ); STEL 100 ppm ACGIH ® ; IDLH = 1000 ppm. Target organ effects from inhalation include fetal, reproductive and behavioral symptoms. Flammable liquid. May overheat dangerously if large quantities are mixed with polymerizing agents. Keep away from strong acids and bases. Common glove materials are not effective; use Teflon. Work in a hood. Polymerize small quantities for disposal.
Nickel chloride. TWA = 1.0 mg nickel/cubic meter (0.1 mg nickel/cubic meter ACGIH ® , 0.015 mg nickel/cubic meter NIOSH); IDLH = 10 mg nickel/cubic meter. Carcinogenic to humans. Toxic by inhalation of dust. Solutions pose little risk to workers but are an environmental problem. Use gloves (any material) and hood when handling the powder. Do not use drain disposal for these solutions or for subsequent rinse fluids.
Nitric acid. TWA = 2 ppm; STEL = 4 ppm ACGIH ® , NIOSH; IDLH = 25 ppm. Corrosive to skin, mucous membranes and most metals. Toxic by inhalation. Target organ effects on reproductive and fetal systems after ingestion. Oxidizer. Concentrated acid is very hazardous. Use Neoprene gloves for extensive use; nitrile, butyl and latex are not effective except to protect against minor splashes. Wear apron and goggles for handling any quantity. Always add acid to water, never water to acid, to avoid severe splattering. Explosive mixtures may be formed with hydrogen peroxide, diethyl ether and anion exchange resins.
Nitrogen, liquid. No PELs established. Asphyxiant gas: excessive inhalation may cause dizziness, unconsciousness or death. Use extreme caution to avoid thermal (cold) burns.
Osmium tetroxide (osmic acid). TWA = 0.0002 ppm osmium; STEL = 0.0006 ppm osmium (ACGIH ® ); IDLH = 0.1 ppm. Vapors are extremely dangerous. Corrosive to eyes and mucous membranes. Toxic by inhalation with target effects on reproductive, sensory and respiratory systems. Avoid all contact with vapors. Do not open containers in air. In a hood, score vial and break under water or other solvent. Information on protective glove materials is not available.
Oxalic acid. TWA = 1 mg/cubic meter; STEL = 2 mg/cubic meter; IDLH = 500 mg/cubic meter. Corrosive solid; causes severe burns of the eyes, skin and mucous membranes. Toxic by inhalation and ingestion, with target organ effects on kidneys and cardiovascular systems. Repeated skin contact can cause dermatitis and slow-healing ulcers. Will corrode most metals. Risks are minimal with quantities usually encountered in histology.
Periodic acid. No PELs established. Mild oxidizer. Quantities used in histology pose little physical or health risk.
Phenol. TWA = 5 ppm; additional exposure likely through skin contact; CL = 15.6 ppm for 15 minutes; IDLH = 250 ppm. Toxic by ingestion, inhalation and skin absorption. Readily absorbed through skin, causing increased heart rate, convulsions and death. Will burn eyes and skin. Target organ effects on digestive, urinary and nervous systems. Combustible liquid (flash point = 172°F). Avoid all contact if possible, or use extreme caution. Purchase the smallest quantity possible. Use only butyl rubber gloves and work only under a fume hood. Mixing concentrated formaldehyde and phenol may produce an uncontrollable reaction.
Phosphomolybdic and phosphotungstic acids. TWA = 1 mg/cubic meter ACGIH ® ; STEL = 3 mg/cubic meter ACGIH ® , NIOSH; IDLH = 1000 mg/cubic meter. All PELs are expressed as the quantity of the metal molybdenum or tungsten. Oxidants. These reagents present minor risk under normal conditions of use in histology.
Picric acid. TWA = 0.1 mg/cubic meter; additional exposure likely through skin contact. Toxic by skin absorption. Explosive when dry or when complexed with metal and metallic salts. Do not move bottles containing dry picric acid; get professional help immediately. Do not allow any picric acid solutions, including yellow rinse fluids or processing solvents, to go down the drain, as these may form explosive picrates with metal pipes. Avoid all use if possible, substituting zinc formalin or glyoxal for Bouin’s or similar fixatives, and tartrazine for a yellow counterstain. If you must have it, check containers monthly to keep the salts wet. Always wipe jar and cap threads with a damp towel to prevent material from drying within them.
Potassium dichromate. See chromic acid for information on chromium toxicity.
Potassium ferricyanide and potassium ferrocyanide. Low toxicity to humans and the environment in quantities likely to be encountered in histology.
Potassium hydroxide. CL = 2 mg/cubic meter as dust (NIOSH, ACGIH ® ). Corrosive to eyes and skin. Use care when dissolving solids in water, as the reaction may be violently exothermic and cause splattering.
Potassium permanganate. Skin and eye irritant. Ingestion will cause severe gastrointestinal distress. Strong oxidant: do not mix with ethylene glycol, ethanol, acetic acid, formaldehyde, glycerol, hydrochloric acid, sulfuric acid, hydrogen peroxide or ammonium hydroxide. Use butyl gloves.
Propidium iodide. Mutagen, irritant and suspected carcinogen. Material is irritating to mucous membranes and upper respiratory tract. All common glove materials except latex are suitable.
Propylene glycol ethers. TWA = 100 ppm; STEL = 150 ppm (ACGIH ® ). Used as a less toxic substitute for ethylene-based glycol ethers.
Pyridine. TWA = 5 ppm (1 ppm ACGIH ® ); IDLH = 1000 ppm. Toxic by ingestion, inhalation and skin absorption. Overexposure causes nausea, headache and increased urinary frequency. Target organ effects on liver and kidneys. Irritant to skin and eyes. Highly offensive odor. Flammable liquid (flash point = 20°C). Use only under a fume hood, with butyl gloves. Do not mix with chromic acid.
Silver salts and solutions. TWA = 0.01 mg silver/cubic meter; IDLH = 10 mg silver/cubic meter. Skin and eye irritants. Ingestion will cause violent gastrointestinal discomfort. Little risk to workers when fresh, but some aged solutions become explosive. Serious environmental hazard. Do not discard solutions or rinse fluids down the drain. Silver may be recoverable in special equipment or by metal reclaimers.
Sodium azide. CL = 0.3 mg/cubic meter for the powder (NIOSH, ACGIH ® ). Poison, very toxic. May be fatal if swallowed or absorbed through the skin. Evolves highly toxic gas when mixed with acids. When used as a preservative in biochemical solutions there is little risk to workers except by ingestion and skin absorption. Forms explosive compounds with metals. Do not discard waste down the drain.
Sodium bisulfite. TWA = 5 mg/cubic meter (NIOSH, ACGIH ® ). Irritant to skin, eyes and mucous membranes. Strong reducing agent: keep from oxidants. Dilute solutions generally pose no risk.
Sodium hydroxide. See potassium hydroxide.
Sodium hypochlorite (liquid chlorine bleach). No PELs established. Eye irritant. May be toxic by ingestion unless diluted considerably. Strong oxidant, corrosive to most metals. All common glove materials provide suitable protection. Do not mix bleach with formaldehyde, aminoethylcarbazole (AEC) or diaminobenzidine (DAB).
Sodium iodate. Little risk likely with laboratory quantities. Use to replace mercuric oxide in Harris hematoxylin.
Sodium metabisulfite. See sodium bisulfite.
Sodium phosphate, monobasic and dibasic. Harmless to workers. May pose an environmental problem from eutrophication (over-enrichment of aquatic systems).
Sodium sulfite. See sodium bisulfite.
Sodium thiosulfate. Health risks are minimal under normal conditions of use in histology. Solutions used to ‘de-Zenkerize’ sections will contain significant amounts of mercury and are not discarded down the drain.
Sulfuric acid. TWA = 1 mg/cubic meter (0.2 mg/cubic meter ACGIH ® ); IDLH = 15 mg/cubic meter. Strong irritant to skin, eyes and respiratory system. Concentrated acid is especially dangerous because it fumes. Target organ effects from inhalation on respiratory, reproductive and fetal systems. Dilute solutions pose little risk. Corrosive to most materials. Use a fume hood, apron, goggles and gloves (any common material except butyl). Always add acid to water, never water to acid, to avoid severe splattering.
Tetrahydrofuran (THF). TWA = 200 ppm; STEL = 250 ppm; IDLH = 2000 ppm; BEI = 50 mg THF/liter urine at end of shift. Toxic by ingestion and inhalation. Vapors cause nausea, dizziness, headache and anesthesia. Liquid can defat the skin. Eye and skin irritant. Flammable liquid. Dangerous fire hazard because of low flash point (5°F) and high evaporation rate. Only Teflon gloves are suitable. Avoid all use, as there is no practical way to protect against skin contact.
Toluene. TWA = 200 ppm (50 ppm ACGIH ® ); STEL = 150 ppm; IDLH = 500 ppm; BEI = 50 mg o -cresol/liter urine at end of shift. Skin and eye irritant. Toxic by ingestion, inhalation and skin contact. Target organ effects on fetal, respiratory and central nervous system. Repeated exposure produces neurotoxic effects (impaired memory, poor coordination, mood swings and permanent nerve damage). Flammable (flash point = 5°C). Avoid all use if possible or restrict use severely. No common glove material will provide adequate protection. Substitute one of the short-chain aliphatic hydrocarbon clearing agents except as a diluent in mounting media and for removing coverslips. Exposure may be monitored by measuring the amount of methylhippuric acids in urine (see page 14)
Trichloroethane. TWA = 350 ppm, STEL = 450 ppm. Irritant to skin and eyes. Target organ effects on gastrointestinal and central nervous systems. Non-combustible. No common glove material is suitable. Chlorinated solvents pose severe environmental risks and serious disposal problems. Avoid all use.
Uranyl nitrate. TWA = 0.05 mg uranium/cubic meter; STEL = 0.6 mg/cubic meter ACGIH ® ; IDLH = 10 mg/cubic meter. Corrosive to tissue and most metals. Highly toxic, with target organ effects on liver, urinary, circulatory and respiratory systems. Radiation hazard from inhalation of fine particles; most substances block radioactivity, so handling solutions poses little risk. Any type of glove material except latex is satisfactory. Severe environmental toxin. Problems with transportation and disposal have made this chemical very difficult or impossible to obtain. Find alternate stains for most uses and employ immunohistochemistry for equivocal cases. This will eliminate both uranyl nitrate and silver from the lab.
Xylene. TWA = 100 ppm; STEL = 150 ppm; IDLH = 900 ppm; BEI = 1.6 g hippuric acid/g creatinine in urine at end of shift. See toluene for further information.
Zinc chloride. Corrosive to most metals, including stainless steel. All common glove materials except latex are satisfactory. Do not use zinc chloride solutions in tissue processors. Skin and eye irritant. Ingestion can cause intoxication and severe gastrointestinal upset.
Zinc formalin. A solution of zinc sulfate or zinc chloride and formaldehyde. See individual entries for those ingredients.
Zinc sulfate. Eye irritant, but otherwise not hazardous in quantities used in histology.

This is the science concerned with the relationship between human beings, the machines and equipment they use and their working environment. It involves the application of physiological, anatomical and psychological data to the design of efficient working systems. In the histology laboratory, ergonomic considerations include work habits, posture, preference for right or left handedness, arrangement and use of instrumentation and tools, countertop heights, seating, lighting, noise levels, temperatures, and vibration. Following a situational analysis, preventive measures can be implemented.
A full discussion of ergonomics is beyond the realms of this text but the main areas pertaining to histopathology are briefly outlined below and in greater detail in the previous edition of this book.
Biomechanical risk factors include exposure to excessive force, repetitive movements, awkward working postures and vibration. A variety of musculoskeletal disorders including tendonitis, tenosynovitis, carpal tunnel syndrome and other nerve disorders can occur if these are not addressed.
The proper balance of work between people and machines is sometimes difficult to determine. Automation is desirable because it reduces the physical stresses imposed on workers, but having too much automation takes away the unique value of human interaction and decision making that is based on visual interpretation and cognitive experience. Automation should be considered to replace manual tasks that require standardization (processing and staining) and those that may contribute to musculoskeletal disorders (slide and cassette labeling, microtomy and coverslipping).
Good workstation design is essential in creating a healthy, comfortable, and task-efficient laboratory. Workstations should follow the workflow and be planned to accommodate all equipment and supplies. Consideration must also be given to the number of people who will use the space, their physical characteristics, whether they will sit, stand, or use a combination of positions, and if they need some type of aid to be able to see and reach all of the necessary components. Air quality, temperature, and humidity must be regulated and drafts must be avoided. Lighting must be task appropriate, not necessarily standard overhead lighting, and noise must be minimal.
Ideally, laboratory workstations are versatile, modular, and flexible so that they can be altered to accommodate new tasks, equipment or people. The work surface should be height adjustable, and seating should be individualized and task appropriate.

Suggestions for specific tasks

Computer operation
• Maintain good posture with joints in a relaxed, neutral position.
• Keep the keyboard at elbow height or tilted downward.
• Use a gentle touch on the keys.
• Do not hold your thumb or little finger in the air.
• Place the mouse by the keyboard. Be aware that the burden is on one hand and finger.
• Position the top of the monitor at eye level.
• Wear glasses (if needed) that allow you to keep your head upright or bent slightly forward.
• Do not cradle the phone on your shoulder while working.
• Eliminate sources of reflections and glare on the monitor screen.
Cassette and slide labeling
• Rest wrists on a padded surface when writing.
• Take rest breaks and vary tasks.
• Avoid excessive reaching.
• Use ergonomic writing utensils with large, padded grips.
• Do not use excessive force.
• Automate if possible.
Changing the solutions on the processor
• Use proper bending and lifting techniques.
• Carry containers using a power grip (whole or both hands).
• Use a stool with safe footing to reach above chest height.
• Maintain good sitting posture.
• Keep as many items as possible within your reach area and use ergonomic tools if available.
• Keep joints in a neutral position. Do not lean arms on sharp or hard surfaces. Take mini-breaks and exercise wrists and fingers.
• Alternate the motion used to open cassette lids.
• Get up periodically and walk around.
Manual microtomy
• Maintain good sitting posture.
• Use a well-adjusted, ergonomic chair and a footrest, if necessary.
• Keep joints in a relaxed, neutral position.
• Do not rock the handwheel (wrist flexion and extension).
• Use a cut-out workstation or an L-shaped extension to reach the water bath without bending at the waist and reaching over.
• Take mini-breaks as often as time constraints allow.
• Automate as soon as possible.
Manual staining
• Maintain good standing posture.
• Prop one foot up or stand with one foot forward, and alternate often.
• Keep work as close to the body as possible.
• Use caution when bending, lifting, and reaching.
• Avoid repeatedly dipping slides (wrist flexion and extension).
• Use slide holders and racks rather than forceps.
• Avoid using excessive force to squeeze bottles.
• Automate as soon as possible.
Manual coverslipping
• Maintain correct posture with head upright and joints in a neutral position.
• Keep work at elbow height and within a close reach.
• Do not lean arms on sharp or hard surfaces.
• Take multiple mini-breaks and do stretching exercises.
• Use ergonomic forceps.
• Maintain correct posture. Work at a cut-out bench if possible.
• Keep work at elbow height and as close as possible.
• Use low-profile tubes, solution containers, and waste receptacles.
• Keep wrists in a neutral position.
• Do not twist or rotate at the waist.
• Use electronic, light-touch pipettes designed for multiple finger use.
• Hold the pipette with a relaxed grip.
• Take short breaks every 20–30 minutes if possible.
• Keep hands as warm as possible to maintain feeling and sensitivity.
• Maintain good posture. Do not lean into the chamber.
• If standing, work with one foot propped up and alternate regularly.
• Keep ancillary items as close as possible (possibly on a cart).
• Use skills detailed under ‘Manual microtomy’.
• Avoid static postures, get up and move around periodically and alternate tasks.
• Work with the head bent slightly down instead of back.
• Use a well-adjusted, ergonomic chair and sit close to the microscope.
• Use armrests with a soft, smooth surface.
• Use a microscope with ergonomically positioned controls.
• Use adjustable eyepieces or mount the microscope at a 30° angle.
• Request extenders for the microscope body if the eyepieces are still not high enough.
• Work in a place away from drafts and noise.


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Centers for Disease Control. Guidelines for protecting the safety and health of health care workers, CDC Publication #88–119 . Washington, DC: US Government Printing Office; 1988.
Centers for Disease Control. Guidelines for preventing the transmission of tuberculosis in health-care settings, with special focus on HIV-related issues. Morbidity and Mortality Weekly Report 39, 1–29 . Washington, DC: US Government Printing Office; 1990.
Centers for Disease Control. Guidelines for preventing the transmission of Mycobacterium tuberculosis in health-care facilities. Morbidity and Mortality Weekly Report . 1994;43:1–132. Washington, DC: US Government Printing Office
Clinical and Laboratory Standards Institute. Clinical laboratory waste management: approved guideline. Document GP05-A2. second ed. Wayne, PA:CLSI; 2002.
Clinical and Laboratory Standards Institute. Clinical laboratory safety: approved guideline. Document GP17-A2, second ed. Wayne, PA:CLSI; 2004.
Clinical and Laboratory Standards Institute. Protection of laboratory workers from occupationally acquired infections: approved guideline. Document M29-MA3, third ed. Wayne, PA:CLSI; 2005.
Dapson J.C., Dapson R.W. Hazardous materials in the histopathology laboratory: regulations, risks, handling and disposal , fourth ed. Battle Creek, MI: Anatech Ltd; 2005.
Esswein E.J., Boeniger M.F. Effect of an ozone-generating air-purifying device on reducing concentrations of formaldehyde in air. Applied Occupational Environmental Hygiene . 1994;9:139–146.
Kiernan J.A. Histological and histochemical methods: theory and practice , third ed. Boston: Butterworth Heinemann; 1999.
Lunn G., Sansone E.B. Destruction of hazardous chemicals in the laboratory . New York: John Wiley; 1990.
Lunn G., Sansone E.B. The safe disposal of diaminobenzidine. Applied Occupational and Environmental Hygiene . 1991;6:49–53.
Montgomery L. Health and safety guidelines for the laboratory . Chicago: American Society of Clinical Pathologists Press; 1995.
National Institute for Occupational Safety and Health, 2003. NIOSH pocket guide to chemical hazards. DHHS (NIOSH) Publication No. 97–140.
National Research Council. Prudent practices for the handling and disposal of infectious materials . Washington, DC: National Academy Press; 1989.
National Research Council. Prudent practices in the laboratory: handling and disposal . Washington, DC: National Academy Press; 1995.
Ontario (Canada) Ministry of Labor. Regulation respecting control of exposure to biological or chemical agents – made under the Occupational Health and Safety Act . Toronto: Ontario Government Publications; 1991.
Rank J.P. How can histotechnologists protect themselves from Creutzfeldt-Jakob disease? Laboratory Medicine . 1999;30:305–306.
Saunders G.T. Laboratory fume hoods: a user’s manual . Cincinnati, Ohio: American Conference of Governmental Industrial Hygienists; 1993.
Schwope A.D., Costas P.P., Jackson J.O., et al. Guidelines for the selection of chemical protective clothing . Cincinnati, Ohio: American Conference of Governmental Industrial Hygienists; 1987.
Stricoff R.S., Walters B.D. Laboratory health and safety handbook: a guide for the preparation of a chemical hygiene plan . New York: Wiley; 1990.
3 Light microscopy

John D. Bancroft, Alton D. Floyd

Light and its properties
Visible light occupies a very narrow portion of the electromagnetic spectrum. The electromagnetic spectrum extends from radio and microwaves all the way to gamma rays. Electromagnetic energy is complex, having properties that are both wave-like and particle-like. A discussion of these topics is well beyond the scope of this chapter. Suffice to say, visible light is that portion of the electromagnetic spectrum that can be detected by the human eye. In physics texts, this range is generally defined as wavelengths of light ranging from approximately 400 nm (deep violet) to 800 nm (far red). Most humans cannot see light of wavelengths much beyond 700 nm (deep red).
It is common practice to illustrate the electromagnetic spectrum as a sine wave. This is a convenient representation as the distance from one sine peak to another represents the wavelength of light ( Fig. 3.1 ). Light that has a single wavelength is monochromatic: that is, a single color. The majority of sources of light provide a complex mixture of light of different wavelengths, and when this mixture approximates the mixture of light that derives from the sun, we perceive this as ‘white’ light. By definition, white light is a mixture of light that contains some percentage of wavelengths from all of the visible portions of the electromagnetic spectrum. It should be understood that almost all light sources provide a mixture of wavelengths of light (exceptions being devices such as lasers, which generate monochromatic, coherent light). One measure of the mixture of light given off by a light source is color temperature. In practical terms, the higher the color temperature, the closer the light is to natural daylight derived from the sun. Natural daylight from the sun is generally stated to have a color temperature of approximately 5200 kelvin (K). Incandescent light, from tungsten bulbs, has a color temperature of approximately 3200K. These values will be familiar to those using color film for photography, as film type must be chosen to suit the illumination source. As a general rule, the higher the color temperature, the more ‘blue’ or white the light appears to the eye. Lower color temperatures appear more red to yellow, and are regarded as being ‘warmer’ in color.

Figure 3.1 Representation of a light ray showing wavelength and amplitude.
Shorter wavelengths of light (toward the blue to violet end of the spectrum) have a higher energy content for a given brightness of light. As one goes to even shorter wavelengths of the electromagnetic spectrum, the energy content becomes even higher (X-rays and gamma rays). The energy content of light is generally expressed as an energy level, or amplitude based on the electron volts per photon (the particle representation of light). Visible light has an energy level of approximately one electron volt per photon, and the energy level increases as one moves toward the violet and ultraviolet range of the spectrum. Approaching the soft X-ray portion of the spectrum, the energy level per photon ranges from 50 to 100 electron volts (eV). It is this higher energy in the shorter wavelengths of light (the ultraviolet and blue end of the spectrum) that is exploited to elicit fluorescence in some materials.
Light sources give off light in all directions, and most light sources consist of a complex mixture of wavelengths. This mixture of wavelengths is what defines the color temperature of the light source. It should also be noted that the mixture of wavelengths is influenced by the type of material making up the source. Since the majority of light sources used in microscopy are either heated filaments or arcs of molten metal, each source will provide a specific set of wavelengths related to the material being heated. This is referred to as the emission spectrum . Some sources provide relatively uniform mixtures of wavelengths, although of different amplitudes or intensities, such as tungsten filament lamps and xenon lamps. Others, such as mercury lamps, provide very discrete wavelengths scattered over a broad range, but with distinct gaps of no emission between these peaks.
Although light sources are inherently non-coherent (with the exception of lasers), standard diagrams of optics always draw light rays as straight lines. This is a simplification, and it should be remembered that the actual light consists of every possible angle of light rays from the source, not just the single ray illustrated in the diagram. Another property of light that is important for an understanding of microscope optics is absorption of some of the light by the medium through which the light passes ( Fig. 3.2 ). This is seen as a reduction in the amplitude, or energy level, of the light. The medium through which the light passes can also have an effect on the actual speed at which the light passes through the material, and this is referred to as retardation .

Figure 3.2 The amplitude (i.e. brightness) diminishes as light gets further from the source because of absorption into the media through which it passes.

Retardation and refraction
Media through which light is able to pass will slow down or retard the speed of the light in proportion to the density of the medium. The higher the density, the greater the degree of retardation . Rays of light entering a sheet of glass at right angles are retarded in speed but their direction is unchanged ( Fig. 3.3a ). If the light enters the glass at any other angle, a deviation of direction will occur in addition to the retardation, and this is called refraction ( Fig. 3.3b ). A curved lens will exhibit both retardation and refraction ( Fig. 3.3c ), the extent of which is governed by:

(a) the angle at which the light strikes the lens – the angle of incidence ,
(b) the density of the glass – its refractive index , and
(c) the curvature of the lens.

Figure 3.3 (a) Rays passing from one medium to another, perpendicular to the interface, are slowed down at the same moment. (b) Rays passing at any other angle to the interface are slowed down in the order that they cross the interface and are deviated. (c) Rays passing through a curved lens exhibit both retardation and refraction.
The angle by which the rays are deviated within the glass or other transparent medium is called the angle of refraction and the ratio of the sine values of the angles of incidence (i) and refraction (r) gives a figure known as the refractive index (RI) of the medium ( Fig. 3.4a ). The greater the RI, the higher the density of the medium. The RI of most transparent substances is known and is of great value in the computation and design of lenses, microscope slides and coverslips, and mounting media. Air has a refractive index of 1.00, water 1.30, and glass a range of values depending on type but averaging 1.50.

Figure 3.4 (a) Angle of incidence (i) and refraction (r). (b) Ray C–D is lost through the edge of the lens. Ray E–F shows total internal reflection. (c) Parallel rays entering a curved lens are brought to a common focus.
As a general rule, light passing from one medium into another of higher density is refracted towards the normal, and when passing into a less dense medium it is refracted away from the normal. The angle of incidence may increase to the point where the light emerges parallel to the surface of the lens. Beyond this angle of incidence, total internal reflection will occur, and no light will pass through ( Fig. 3.4b ).

Image formation
Parallel rays of light entering a simple lens are brought together by refraction to a single point, the ‘principal focus’ or focal point , where a clear image will be formed of an object ( Fig. 3.4c ). The distance between the optical center of the lens and the principal focus is the focal length . In addition to the principal focus, a lens also has other pairs of points, one either side of the lens, called conjugate foci , such that an object placed at one will form a clear image on a screen placed at the other. The conjugate foci vary in position, and as the object is moved nearer the lens the image will be formed further away, at a greater magnification, and inverted. This is the ‘ real image ’ and is that formed by the objective lens of the microscope ( Fig. 3.5 ).

Figure 3.5 A real image is formed by rays passing through the lens from the object, and can be focused on a screen.
If the object is placed yet nearer the lens, within the principal focus, the image is formed on the same side as the object, is enlarged, the right way up, and cannot be projected onto a screen. This is the ‘ virtual image ’ ( Fig. 3.6 ), and is that formed by the eyepiece of the microscope of the real image projected from the objective. This appears to be at a distance of approximately 25 cm from the eye – around the object stage level. Figure 3.7 illustrates the formation of both images in the upright compound microscope, as is commonly used in histopathology.

Figure 3.6 A virtual image is viewed through the lens. It appears to be on the object side of the lens.

Figure 3.7 Ray path through the microscope.
The eye sees the magnified virtual image of the real image, produced by the objective.

Image quality
White light is composed of all spectral colors and, on passing through a simple lens, each wavelength will be refracted to a different extent, with blue being brought to a shorter focus than red. This lens defect is chromatic aberration ( Fig. 3.8a ) and results in an unsharp image with colored fringes. It is possible to construct compound lenses of different glass elements to correct this fault. An achromat is corrected for two colors, blue and red, producing a secondary spectrum of yellow/green, which in turn can be corrected by adding more lens components – the more expensive apochromat .

Figure 3.8 (a) Chromatic aberration. (b) Spherical aberration.
Microscope objectives of both achromatic and apochromatic types (see Fig. 3.11 ) are usually overcorrected for longitudinal chromatic aberration and must be combined with matched compensating eyepieces to form a good-quality image. This restriction on changing lens combinations is overcome by using chromatic, aberration-free (CF) optics, which correct for both longitudinal and lateral chromatic aberrations and remove all color fringes, being particularly useful for fluorescence and interference microscopes.
Other distortions in the image may be due to coma, astigmatism, curvature of field, and spherical aberration, and are due to lens shape and quality. Spherical aberration is caused when light rays entering a curved lens at its periphery are refracted more than those rays entering the center of the lens and are thus not brought to a common focus ( Fig. 3.8b ).
These faults are also corrected by making combinations of lens elements of different glass, e.g. fluorite, and of differing shapes.

The components of a microscope

Light source
Light, of course, is an essential part of the system; at one time sunlight was the usual source. A progression of light sources has developed, from oil lamps to the low-voltage electric lamps of today. These operate via a transformer and can be adjusted to the intensity required. The larger instruments have their light sources built into them. Dispersal of heat, collection of the greatest amount of light, and direction and distance are all carefully calculated by the designer for greatest efficiency. To obtain a more balanced white light approximation, these light sources must often be operated at excessive brightness levels. The excess brightness is reduced to comfortable viewing levels through the use of neutral density filters .

Light from the lamp is directed into the first major optical component, the substage condenser, either directly or by a mirror or prism. The main purpose of the condenser is to focus or concentrate the available light into the plane of the object ( Fig. 3.9 ). Within comfortable limits, the more light at the specimen, the better is the resolution of the image.

Figure 3.9 The function of the condenser is to concentrate, or focus, the light rays at the plane of the object.
Many microscopes have condensers capable of vertical adjustment, in order to allow for varying heights or thickness of slides. Once the correct position of the condenser has been established, there is no reason to move it, as any alteration will change the light intensity and impair the resolution. In most cases condensers are provided with adjustment screws for centering the light path. Checking and, if necessary, adjusting the centration before using the instrument should be a routine procedure for every microscopist. All condensers have an aperture diaphragm with which the diameter of the light beam can be controlled.
Adjustment of this iris diaphragm will alter the size and volume of the cone of light focused on the object. If the diaphragm is closed too much, the image becomes too contrasty and refractile, whereas if the diaphragm is left wide open, the image will suffer from glare due to extraneous light interference. In both cases the resolution of the image is poor. The correct setting for the diaphragm is when the numerical aperture of the condenser is matched to the numerical aperture of the objective in use ( Fig. 3.10 ) and the necessary adjustment should be made when changing from one objective to another. This is achieved, approximately, by removing the eyepiece, viewing the substage iris diaphragm in the back focal plane of the objective, and closing it down to two-thirds of the field of view.

Figure 3.10 Rays A illustrate the ‘glare’ position resulting in extraneous light and poor resolution. Rays B indicate the correct setting of the substage iris diaphragm.
With experience the correct setting can be estimated from the image quality. Under no circumstances should the iris diaphragm be closed to reduce the intensity of the light; use filters or the rheostat of the lamp transformer. Many condensers are fitted with a swing-out top lens. This is turned into the light path when the higher-power objectives are in use. It focuses the light into a field more suited to the smaller diameter of the objective front lens. Swing it out of the path with the lower power objectives, or the field of view will only be illuminated at the center. When using apochromatic or fluorite objectives the substage condenser should also be of a suitable quality, such as an aplanatic or a highly corrected achromatic condenser.

Object stage
Above the condenser is the object stage, which is a rigid platform with an aperture through which the light may pass. The stage supports the glass slide bearing the specimen, and should therefore be sturdy and perpendicular to the optical path. In order to hold the slide firmly, and to allow the operator to move it easily and smoothly, a mechanical stage is either attached or built in. This allows controlled movement in two directions, and in most cases Vernier scales are incorporated to enable the operator to return to an exact location in the specimen at a later occasion.

The next and most important piece of the microscope’s equipment is the objective, the type and quality of the objective having the greatest influence on the performance of the microscope as a whole.
Within the objective there may be from 5 to 15 lenses and elements, depending on image ratio, type and quality ( Fig. 3.11 ). The main task of the objective is to collect the maximum amount of light possible from the object, unite it, and form a high-quality magnified real image, some distance above. Older microscopes used objectives computed for an optical tube length of 160 mm (DIN standard), or 170 mm (Leitz only), but these fixed tube length systems have now been largely replaced by infinity corrected objectives that can greatly extend this tube length and permit the addition of other devices into the optical path.

Figure 3.11 Diagram of achromatic and apochromatic objectives.
Some examples of the latter may have as many as 15 separate lens elements.
Magnifying powers or, more correctly, object-to-image ratios of objectives are from 1 : 1 to 100 : 1 in normal biological instruments.
The ability of an objective to resolve detail is indicated by its numerical aperture and not by its magnifying power. The numerical aperture or NA is expressed as a value, and will be found engraved on the body of the objective. The value expresses the product of two factors and can be calculated from the formula:

where n is the refractive index of the medium between the coverglass over the object and the front lens of the objective, for example air, water, or immersion oil, and u is the angle included between the optical axis of the lens and the outermost ray that can enter the front lens ( Fig. 3.12 ).

Figure 3.12 The refractive index of the medium between the coverglass and the surface of the objective’s front lens (in this case air, RI = 1.00), and the sine of the angle ( u ) between the optical axis and the outermost accepted ray (r), gives the numerical aperture (see text).
In Figure 3.12 the point where the axis meets the specimen is regarded as a light source; rays radiate from this point in all directions. Some will escape to the outside, and some will be reflected back from the surface of the coverglass. Ray r is the outermost ray that can enter the front lens; the angle u between ray r and the axis gives us the sin value required. In theory the greatest possible angle would be if the surface of the front lens coincided with the specimen, giving a value for u of 908. In the above formula, with air (RI = 1.00) as the medium, and a value for u of 908 ( sin u × 1), the resulting NA = 1.00. Of course this is impossible as there must always be some space between the surfaces, so a value of 908 for u is unobtainable. In practice the maximum NA attainable with a dry objective is 0.95. Similar limitations apply to water and oil immersion objectives; theoretical maximum values for NA are 1.30 and 1.50, respectively. In practice values of 1.20 and 1.40 are the highest obtainable.
Resolution does not depend entirely on the NA of a lens but also on the wavelength of light used, governed by the following relationship:

where the resolution is the smallest distance between two dots or lines that can be seen as separate entities, and λ is the wavelength of light.
The resolving power of the objective is its ability to resolve the detail that can be measured. In summary, as the NA of an objective increases, the resolving power increases but working distance, flatness of field, and focal length decrease.
Objectives are available in varying quality and types ( Fig. 3.11 ). The achromatic is the most widely used for routine purposes; the more highly corrected apochromats , often incorporating fluorite glass, are used for more critical work, while plan-apochromats (which have a field of view that is almost perfectly flat) are recommended for photomicrography. For cytology screening, flat-field objectives – often plan-achromats – are particularly useful. On modern microscopes, up to six objectives are mounted onto a revolving nosepiece to enable rapid change from one to another and, ideally, the focus and field location should require the minimum of adjustment. Such lenses are said to be par-focal and par-central .
Most objectives are designed for use with a coverglass protecting the object. If so, a value giving the correct coverglass thickness should be found engraved on the objective. Usually this is 0.17 mm. Some objectives, notably apochromats between 40 : 1 and 63 : 1, require coverslip thickness to be precise. Some are mounted in a correction mount and can be adjusted to suit the actual thickness of the coverglass used.

Body tube
Above the nosepiece is the body tube. Three main forms are available: monocular, binocular, and the combined photo-binocular. The last sometimes has a prism system allowing 100% of the light to go either to the observation eyepieces, or to the camera located on the vertical part, and sometimes has a beam-splitting prism dividing the light, 20% to the eyes and 80% to the camera. This facilitates continuous observation during photography. Provision is made in binocular tubes for the adjustment of the interpupillary distance, enabling each observer to adjust for the individual facial proportions. Alteration of this interpupillary distance may alter the mechanical tube length, and thus the length of the optical path. This can be corrected either by adjusting the individual eyepiece tubes, or by a compensating mechanism built into the body tube.
Modern design tends towards shortening the physical lengths of the components, and in consequence, the intermediate optics are sometimes included in the optical path to compensate. These lenses are mounted on a rotating turret and are designated by their magnification factor. Additionally, a tube lens may be incorporated for objectives that are infinity corrected , as these objectives form only a virtual image of the object, which must be converted to a real image focused at the lower focal plane of the eyepiece.

Eyepieces are the final stage in the optical path of the microscope. Their function is to magnify the image formed by the objective within the body tube, and present the eye with a virtual image, apparently in the plane of the object being observed; usually this is an optical distance of 250 mm from the eye.
Early types of eyepiece, like objectives, were subject to aberrations, especially of color. Compensating eyepieces were designed to overcome these problems and can be used with all modern objectives. The eyepiece designed by Huyghens in 1690 is still available, together with periplanatic (flat-field) and wide-field types, and eyepieces for holding measuring graticules and photographic formats. High focal point eyepieces are designed for spectacle wearers. For older fixed tube length microscopes, manufacturers often placed different amounts of the various corrections in the optical train in either the objective or the eyepiece. Therefore it is important to use eyepieces from the same manufacturer with objectives from that manufacturer. Eyepieces designed for infinity objectives must be used with the newer infinity-corrected systems.

Magnification and illumination

Magnification values
Total magnification is the product of the magnification values of the objective and eyepiece, provided the system is standardized to an optical tube length of 160 mm. For variations of the latter the formula is:

Where additional tube lenses are included, simply multiply by the designated factor; for example, objective 40x, eyepiece 10x, and tube lens factor 1.25x gives a total magnification of 500x. Choosing the correct eyepiece magnification is important, as a total magnification may be reached without further resolution of the object; this is empty magnification . As a guide, total magnification should not exceed 1000 × NA of the objective. Therefore an objective designated 100/1.30 would allow a total magnification of 1300 (1000 × 1.3NA), so eyepieces in excess of 12.5x would serve no useful purpose. For accurate measurements, calibration of the optics with a stage micrometer is necessary.

Critical illumination , often used with simple equipment and a separate light source, is when the light source is focused by the substage condenser in the same plane as the object, when the object is in focus ( Fig. 3.13 ). At one time, ribbon filament lamps were available for microscope illumination. Modern filament lamps use a spring-like filament, and the image of the filament causes uneven illumination, which is unacceptable.

Figure 3.13 (a) Critical illumination. (b) Köhler illumination.
For photography and all the specialized forms of microscopy it is best to use Köhler illumination , where an image of the light source is focused by the lamp collector or field lens in the focal plane of the substage condenser (on the aperture diaphragm).
The image of the field or lamp diaphragm will now be focused in the object plane and the illumination is even. The image of the light source and the aperture diaphragm will in turn be focused at the back focal plane of the objective and can be examined with the eyepiece removed. Poor resolution will result unless the illumination is centered with respect to the optical axis of the microscope. Figure 3.13 shows the main differences between critical and Köhler systems.

Dark-field illumination
So far the microscope has been shown as suitable for the examination of stained preparations. Staining aids the formation of images by absorbing part of the light (some of the wavelengths) and producing an image of amplitude differences and color. Occasions arise when it is preferable, or essential, that unstained sections or living cells are examined. Such specimens and their components have refractive indices close to that of the medium in which they are suspended and are thus difficult to see by bright-field techniques, due to their lack of contrast. Dark-field microscopy overcomes these problems by preventing direct light from entering the front of the objective and the only light gathered is that reflected or diffracted by structures within the specimen ( Fig. 3.14 ). This causes the specimen to appear as a bright image on a dark background, the contrast being reversed and increased. Dark field permits the detection of particles smaller than the optical resolution that would be obtained in bright field, due to the high contrast of the scattered light. In the microscope, oblique light is created by using a modified or special condenser that forms a hollow cone of direct light which will pass through the specimen but outside the objective ( Fig. 3.14 ). Dark field condensers may be for either dry, low-power objectives or for oil immersion high-power objectives. Whichever is used, the objective must have a lower numerical aperture than the condenser (in bright-field illumination, optimum efficiency is obtained when the NAs of both objective and condenser are matched). In order to obtain this condition it is sometimes necessary to use objectives with a built-in iris diaphragm or, more simply, by inserting a funnel stop into the objective. Perfect centering of the condenser is essential, and with the oil immersion systems it is necessary to put oil between the condenser and the object slide in addition to the oil between the slide and the objective. As only light diffracted by the specimen will enter the objective, a high-intensity light source is required.

Figure 3.14 In dark-field illumination no direct rays enter the objective. Only scattered rays from the edges of structures in the specimen form the image (dashed lines).
Most bright-field microscopes can be converted for dark-field work by using simple patch stops, made of black paper, placed on top of the condenser lens or suspended in the filter holder. Alternatively the patch stops can be constructed from different colored filters ( Rheinberg illumination ) using a dark color for the center disc and a contrasting lighter color for the periphery. This system reduces the glare of conventional dark field and reveals the specimen in, say, red on a blue background.
Variable intensity dark field is obtained by making the Rheinberg discs from polarizing filters, the center being oriented at right angles to the periphery. This allows good photomicrography. Dark-field illumination is particularly useful for spirochetes, flagellates, cell suspensions, flow cell techniques, parasites, and autoradiographic grain counting, and was once commonly used in fluorescence microscopy. Thin slides and coverglasses should be used and the preparation must be free of hairs, dirt, and bubbles. Many small structures are more easily visualized by dark-field techniques due to increased contrast, although resolution may be inferior to bright-field microscopy.

Phase contrast microscopy
Unstained and living biological specimens have little contrast with their surrounding medium, even though small differences of refractive index (RI) exist in their structures. To see them clearly involves either:

a. closing down the iris diaphragm of the condenser, which reduces its numerical aperture (NA) producing diffraction effects and destroying the resolving power of the objective, or
b. using dark-field illumination, which enhances contrast by reversal, but often fails to reveal internal detail.
Phase contrast overcomes these problems by a controlled illumination using the full aperture of the condenser and improving resolution. The higher the RI of a structure, the darker it will appear against a light background, i.e. with more contrast.

Optical principle
If a diffraction grating is examined under the microscope, diffraction spectra are formed in the back focal plane (BFP) of the objective due to interference between the direct and diffracted rays of light. The grating consists of alternate strips of material with slightly different RIs, through which light acquires small phase differences, and these form the image. Unstained cells are similar to diffraction gratings as their contents also differ very slightly in RI.
Two rays of light from the same source with the same frequency are said to be coherent, and when recombined they will double in amplitude or brightness if they are in phase with each other ( constructive interference ). However, if they are out of phase with each other, destructive interference will occur.
Figure 3.15a represents the waveform of a light ray. In Figure 3.15b the rays are identical but one is out of phase with the other and they interfere but with no increase in amplitude. Figure 3.15c shows one ray now out of phase with the other, and they cancel each other out. This is maximum destructive interference and no light is seen, resulting in maximum contrast. However, if one ray is brighter than the other (increased amplitude) but is still out of phase ( Fig. 3.15d ) then the difference in amplitude can be seen, while maintaining maximum interference. This last position is that which occurs in the phase contrast microscope.

Figure 3.15 Interference of light rays in phase contrast microscopy.

The phase contrast microscope
To achieve phase contrast the microscope requires modified objectives and condenser, and relies on the specimen retarding light by between and . An intense light source is required to be set up for Köhler illumination.
The microscope condenser usually carries a series of annular diaphragms made of opaque glass, with a clear narrow ring, to produce a controlled hollow cone of light. Each objective requires a different size of annulus, an image of which is formed by the condenser in the back focal plane (BFP) of the objective as a bright ring of light ( Fig. 3.16 ). The objective is modified by a phase plate which is placed at its BFP ( Fig. 3.16 ). A positive phase plate consists of a clear glass disc with a circular trough etched in it, to half the depth of the disc. The light passing through the trough has a phase difference of compared to the rest of the plate. The trough also contains a neutral-density light-absorbing material to reduce the brightness of the direct rays, which would otherwise obscure the contrast obtained.

Figure 3.16 A = annulus at focal plane of condenser; B = object plane; C = phase plate at BFP of objective; D = light rays diffracted and retarded by specimen, total retardation compared with direct light; E = direct light rays unaffected by specimen.
It is essential that the image of the bright annular ring from the condenser is centered and superimposed on the dull trough of the objective phase plate. This is achieved by using either a focusing telescope in place of the eyepiece or a Bertrand lens situated in the body tube of the microscope. Each combination of annulus and objective phase plate will require centering. When the hollow cone of direct light from the annulus enters the specimen, some will pass through unaltered while some rays will be retarded (or diffracted) by approximately . The direct light will mostly pass through the trough in the phase plate while the diffracted rays pass through the thicker clear glass and are further retarded.
The total retardation of the diffracted rays is now and interference will occur when they are recombined with the direct light. Thus an image of contrast is achieved, revealing even small details within unstained cells. This is a quick and efficient way of examining unstained paraffin, resin, and frozen sections, as well as studying living cells and their behavior.

Interference microscopy
In phase contrast microscopy, the specimen retards some light rays with respect to those which pass through the surrounding medium. The resulting interference of these rays provides image contrast but with an artifact called the ‘phase halo’. In the interference microscope the retarded rays are entirely separated from the direct or reference rays, allowing improved image contrast, color graduation, and quantitative measurements of phase change (or ‘optical path difference’), refractive index, dry mass of cells (optical weighing), and section thickness.
Whenever light passes across the edge of an opaque object the rays close to that edge are diffracted, or bent away from their normal path. If, instead of a single edge, the rays pass through a narrow slit, then the rays at the edge of the beam will fan out on either side to quite wide angles ( Fig. 3.17a ). Two slits closely side by side form two fans of rays which will cross ( Fig. 3.17b ) and, if coherent, will observably ‘interfere’. If each ray is regarded as a wave it can be seen that phase conditions of increased amplitude and extinction are bound to occur at points where the waves cross and interfere ( Figs 3.17c, d ). The result of this in the microscope is a series of parallel bands, alternately bright and dark across the field of view. With white light, bands of the spectral colors are seen, because the wavelengths making up white light are diffracted at different angles. With monochromatic light, the bands are alternately dark and light, and of a single color. The same effect can be shown if separate beams of coherent light are reunited. This phenomenon is known as ‘interference’.

Figure 3.17 Diffraction and interference of coherent rays (see text).
Early microscope models split a light beam into two parts, each traversing two sets of perfectly matched optics, one beam passing through the specimen (measuring beam) and the other acting as a reference beam. The beams were widely separated and suitable only for large specimens and interference fringe measurements. Later models used a double-beam system, where the separation is produced by birefringent materials and is close enough to require only one objective ( Fig. 3.18a ).

Figure 3.18 (a) Ray path of an interference microscope using a single objective. The beams should be separated sufficiently for one to pass through an empty part of the preparation, otherwise the ‘ghost’ images formed can cause confusion. (b) Appearance of interference bands in the field of view.
If the two paths are equal and in the same phase, the interference bands can be seen running straight and parallel across the field. If an object is introduced into one beam path that causes some shift in the phase, this will be seen as a displacement in the interference bands ( Fig. 3.18b ). When using monochromatic light, each interval comprising one dark and one light band is one wavelength wide, and thus the distance in nanometers is known. Displacement of the bands is measured with a micrometer eyepiece and with this information, coupled with either the RI or object thickness, the measurements referred to earlier can be determined.
Two types of double-beam system have been used. One involved focusing the reference beam below the object – the ‘double focus’ system – and the other involved a lateral displacement of the reference beam called ‘shearing’, where the separation of the beams is very small. Figure 3.19a , b illustrates this latter system using polarized light and Wollaston prisms. The first birefringent prism in the condenser separates the beams and after passing through the object they are recombined by the second identical prism at the back of the objective. A different pair of prisms is required for each magnification. This produces ‘interference contrast’ and together with rotation of the polarizers enhances the three-dimensional (3D) effect in the image. In 1952, Nomarski modified the Wollaston prisms, so that the lateral separation is less than the resolving power of the microscope, producing excellent 3D colored images from unstained specimens. This system is referred to as differential interference contrast or DIC. Additionally, only one such prism is required at the objective level for all magnifications. This system permits enhanced visualization of immunohistochemical preparations.

Figure 3.19 (a) A Wollaston prism is so constructed that rays passing through the center are in phase. Those passing at other points have a phase difference. The arrows and dot represent the optic axes of the prisms, being at right angles to each other. (b) Ray path in the microscope. Each ray is polarized on separation and they vibrate at right angles to each other, producing interference colors when recombined.

Polarized light microscopy
The use of polarized light in microscopy has many useful and diagnostic applications. Numerous crystals, fibrous structures (both natural and artificial), pigments, lipids, proteins, bone, and amyloid deposits exhibit birefringence. Every cellular pathology laboratory should have at least a simple system of polarizing microscopy.
Earlier in this chapter, light was described as a series of pulses of energy radiating away from a source, and shown diagrammatically as a sine curve, with wavelength and amplitude defined. Light can also be described as an electromagnetic vibration, which travels outwards from the source of its propagation, much in the same way as a vibration will travel along a rope when it is jerked in a direction at right angles to its length. The vibrations in the rope will be generated in the direction of the force that caused them, and this is called the plane of vibration , or vibration direction ( Fig. 3.20 ). Natural light vibrates in many planes or vibration directions, whereas polarized light vibrates in only one plane, as in the rope, and can be produced for microscopy purpose by passing natural light through a polarizer, which is an optical component made from a substance that will allow vibrations of only one vibration direction to pass.

Figure 3.20 Plane of vibration produced on a rope.
Substances or crystals capable of producing plane-polarized light are called birefringent . Light entering a birefringent crystal such as calcite is split into two light paths, each determined by a different refractive index (RI) and each vibrating in one direction only (i.e. polarized) but at right angles to each other ( Fig. 3.21 ). The higher the RI, the greater the retardation of the ray, so that each ray leaves the crystal at a different velocity. The high RI ray is called slow and the low RI ray is called fast . There is also a phase difference between the rays, so that, if they are recombined, interference occurs and various spectral colors are seen.

Figure 3.21 A birefringent crystal such as calcite can split a ray of light into two light paths, each vibrating at right angles to the other.
Originally, polarizers, made from calcite and known as Nicol prisms after their inventor, were cemented together with Canada balsam in such a way that the slow ray was reflected away from the optical path and into the mount of the prism, leaving only the polarized fast ray to pass through ( Fig. 3.22 ).

Figure 3.22 A Nicol prism is constructed so that one part of the ray is allowed to pass whilst the other is directed away from the optical path and is lost.
There will be a direction within a birefringent crystal along which light may pass unaltered; this is called the optic axis (see Wollaston prism). Substances through which light can pass in any direction and at the same velocity are called isotropic and are not able to produce polarized light. A knowledge of RI and polarization measurements identifies many crystalline structures and is particularly useful to the materials scientist but is of limited use to the histologist.
Some substances and crystals can produce plane polarized light by differential absorption and give rise to the phenomenon of dichroism . Such crystals suspended in thin plastic films and oriented in one direction have replaced the bulky and expensive Nicol prisms. These thin films totally absorb the slow rays and are pleochroic (absorbing all colors equally), and are the most useful in microscopy as they occupy very little space and can be used with any microscope.
The dedicated polarizing microscope uses two polarizers ( Fig. 3.23 ). One, always referred to as the polarizer , is placed beneath the substage condenser and held in a rotatable graduated mount, and can be removed from the light path when not required. The other, called the analyzer , is placed between the objective and the eyepiece and is also graduated for measurement to be taken. A circular rotating stage would also be present for rotation of the specimen.

Figure 3.23 A microscope equipped for polarized light.
The human eye is not able to distinguish any difference between polarized and natural light, although when looking through a single polarizer there is an obvious loss of intensity, some of which is due to the color of the filter, as well as the splitting and absorption of the rays. Polarizing spectacles used as sunglasses make full use of both properties, but their chief advantage is the elimination of glare and reflected light from such surfaces as water and glass, which act as polarizers, much of the reflected light being polarized at right angles to that which penetrates the surface. Looking through two polarizers, if their vibration directions are parallel, results in a further slight loss of intensity ( Fig. 3.24a ), due to the increase of the thickness and subsequent absorption, but as one is rotated in relation to the other, intensity decreases to extinction when the vibration directions are crossed, and at right angles ( Fig. 3.24b ). The first polarizer only allows the passage of rays vibrating in its own vibration direction; if parallel, the second polarizer will allow those rays to pass; if crossed, passage of the rays is blocked.

Figure 3.24 (a) When polarizer and analyzer are parallel, rays vibrating in the parallel plane are able to pass. (b) When polarizer and analyzer are crossed, rays able to pass the polarizer are blocked by the analyzer. The condition when no light reaches the observer is known as extinction .
Two phenomena detected in polarized light are interesting to the histologist: birefringence and dichroism. When a birefringent substance is rotated between two crossed polarizers, the image appears and disappears alternately at each 45° of rotation. Hence, in a complete revolution of 360° the image appears four times, and likewise, it is extinguished completely four times. In a thin section of rock composed of many types of crystal this phenomenon is very dramatic, especially when interference colors, due to varying thickness of crystal, are present. When one of the planes of vibration of the object is in a parallel plane to the polarizer, only one part ray can develop, and its further passage is blocked by the analyzer in the crossed position. At 45°, however, phase differences between the two rays which can develop are able to combine in the analyzer and form a visible image ( Fig. 3.25 ).

Figure 3.25 When a birefringent substance is rotated between crossed polarizers, it is visible when it is in the diagonal position (i.e. when it is halfway (45°) between the vibration planes of the polarizers). Extinction occurs when one of its planes of vibration is parallel to either polarizer. Both conditions occur four times in a complete revolution of 360°.
Some birefringent substances are also dichroic, which is the second of the phenomena useful to the histologist. Only the polarizer is used and, if no rotating stage is available, the polarizer itself can be rotated. Changes in intensity and color are seen during rotation. The color changes in a rotation of 90°, and back to its original color in the next 90° ( Fig. 3.26 ). This is due to differential absorption of light, depending upon the vibration direction of the two rays in a birefringent substance.

Figure 3.26 A dichroic substance rotated in polarized light (i.e. using polarizer only). Changes of color and intensity can be seen after rotating 90°. The original color returns after a further 90° rotation. This is due to the differential absorption of the two rays in some birefringent substances, depending on the direction of the polarization.
Weak birefringence in biological specimens is enhanced by the addition of dyes or impregnating metals, in an orderly linear alignment, for example along amyloid fibrils. Although only one polarizer is needed to detect the resulting dichroism, the use of the analyzer in addition can enhance the image.

Sign of birefringence
Reference was made earlier to the separated slow and fast rays in a birefringent substance. Additionally, if the slow ray (higher RI) is parallel to the length of the crystal or fiber, the birefringence is positive . If the slow ray is perpendicular to the long axis of the structure, the birefringence is negative . The sign of birefringence is diagnostically useful and is determined by the use of a compensator (birefringent plate of known retardation) either above the specimen or below the polarizer at 45° to the direction of polarized light. Rotate the compensator or the specimen until the slow direction of the compensator (indicated by arrows) is parallel to the long axis of the crystal or fiber. The field is now red and if the crystal is blue the birefringence is positive . If the crystal is yellow , the slow direction of the compensator is parallel to the fast direction of the crystal and the birefringence is negative . Quartz and collagen exhibit positive birefringence while polaroid discs, calcite, urates, and chromosomes are negative. Simple compensators can be made from mica or layers of sellotape.

Fluorescence microscopy
Fluorescence is the property of some substances which, when illuminated by light of a certain wavelength, will re-emit the light at a longer wavelength. In fluorescence microscopy, the exciting radiation is usually in the ultraviolet wavelength (c. 360 nm) or blue region (c. 400 nm), although longer wavelengths can be used with some modern dyes.
A substance that possesses a ‘fluorophore’ will fluoresce naturally. This is known as primary fluorescence or autofluorescence . Ultraviolet excitation is required for optimum results with substances such as vitamin A, porphyrins, and chlorophyll. Dyes, chemicals, and antibiotics added to tissues produce secondary fluorescence of structures and are called ‘fluorochromes’. This is the most common use of fluorescence microscopy and the majority of fluorochromes require only blue light excitation.
Induced fluorescence is a term applied to substances such as the catecholamines, which after treatment with formaldehyde vapor are converted to fluorescent quinoline compounds.
The applications of fluorescence microscopy are numerous in both qualitative and quantitative systems, and some of these are contained in other chapters of this book.

Transmitted light fluorescence

Light sources
All light sources emit a wide range of wavelengths, including the shorter ultraviolet and blue wavelengths which are of interest for fluorescence. Only a few sources emit sufficient short-wave light for practical use. The most commonly used are high-pressure gas lamps, such as the mercury vapor and xenon gas lamps. For some wavelength excitation, in the blue and green range for instance, halogen filament lamps produce enough light to be useful. The choice of a suitable source depends upon the type of work to be performed, and for routine observation purposes it is better to use the mercury vapor burners. These operate on alternating current and their starting equipment is not so costly. The xenon burners operate on direct current and require rectifiers to be included with the starter equipment if they are to be used on normal mains supply. Xenon burners on a DC supply can be stabilized and are therefore suitable for fluorimetry or the measurement of fluorescence emission. The two types of lamp differ in their emission curves: that is to say, mercury lamps reach very high amplitudes at some wavelengths, whereas at other parts of the wavelength range the emission is low, and the curve in general, has a very spiky profile. Xenon, on the other hand, has a smoother, more continuous curve. Fortunately the peaks in the mercury vapor emission coincide with the excitation wavelengths of some of the more widely used fluorochromes.
Because they contain gas at high pressure, these burners must be handled with great care, and must be housed in strong, protective lamphouses. Heat and infrared waves are filtered out before the light from the source begins its journey. At one time all fluorescence systems used the transmitted light route common to normal light microscopy. However, nowadays, the incident route is widely used. High-pressure arc lamps also have specific lifetimes, and in the case of mercury lamps this is only 200 hours. Operation for longer times than this may result in explosive destruction of the lamp, with release of mercury vapor into the immediate vicinity.
In recent years, a new type of illumination source has been introduced. This is the LED source, which is a type of solid-state, semiconductor device. These LED sources have exceptionally long lifetimes, with little change in light output over that lifetime. Another characteristic is that these sources emit light of a single wavelength, with a very narrow peak width. These advantageous features mean that these devices are finding increasing use in illuminators for microscopy. While they are not yet as bright as arc lamps, they do not require much optical filtering to select the wavelength of interest, and they do not suffer from the flicker and ‘arc wandering’ of the high-pressure lamps. They also do not pose the risk of explosion and they have essentially unlimited lifetimes. For different emission wavelengths, different LEDs are required, but there is no need to select a new set of emission filters as the LED itself provides a narrow band of excitation light.

Preparations for fluorescence may contain other fluorescing material in addition to that in which one is interested. It is necessary therefore to filter out all but the specific excitation wavelength to avoid confusion between the important and the unimportant fluorescence ( Fig. 3.27 ).

Figure 3.27 Light path for transmitted fluorescence. Light of all wavelengths passes from the source through a heat-absorbing filter, into a second filter which removes red light, and then through an exciter filter, which allows only the desired wavelength(s) to pass. On passing through the specimen, the objective collects both exciting and fluorescent wavelengths. The former is removed by a barrier filter to protect the eye of the observer.
A variety of filters are available for this purpose. ‘Dyed in the glass’ filters, with such designations as UG 1 and BG 12, are broadband filters and transmit a wide range of wavelengths, the width of the range depending upon the composition and thickness of the filter. Besides the possibility of non-specific and autofluorescence, there may also be materials that are excited at more than one excitation wavelength, so it is better to employ filters of a narrower band transmission that have their transmission peaks closer to the excitation maximum of the fluorochrome, such as FITC. Narrow band filters are often of the ‘interference’ filter type, and are vacuum-coated layers of metals on a glass support. They have a mirror-like surface, and must be inserted in the beam with the reflective face towards the light source. The better-quality filters are carefully selected for their transmission characteristics, and only a few are finally judged suitable. For this reason, they are expensive; careful handling to avoid corrosive finger marks and scratches is essential. When used with high-intensity light sources, these filters may also degrade with time, and so should be checked with an accurate spectrophotometer at regular intervals.
Barrier or suppression filters are placed before the eyepiece to prevent short wavelength light from damaging the retina of the eye ( Fig. 3.27 ). They must, however, allow the fluorescing color to pass; otherwise a negative result may be obtained. Barrier filters are colorless through yellow to dark orange and of specific wavelength transmission. For example a K.470 filter will block all wavelengths below 470 nm. Colored barrier filters may alter the final color rendering of the fluorescent specimen, and for this reason all filters used in the system must be recorded when reporting results.

Condensers for fluorescence microscopy
Bright-field condensers are able to illuminate the object, using all the available energy, but they also direct the rays beyond the object into the objective. Not only is this a potential hazard to the eyes of the observer but also it can set up disturbing autofluorescence in the cement and component layers in the objective itself. In consequence most systems employ a dark-field condenser which does not allow direct light into the objective, and in addition is more certain to give a dark contrasting background to the fluorescence. At the same time it should be realized that only about one-tenth of the available energy is used, limited by the design of the condenser.
Fluorescent light emission is in most cases very poor in relation to the amount of energy absorbed by fluorochromes or fluorophores, with an efficiency ratio somewhere between 1 : 1000 and 1 : 100 at best, and so any system that reduces the available energy to any extent should be well considered before being put into use.

Objectives, too, must be carefully chosen. It has already been noted that autofluorescence is a hazard with bright-field illumination, and for that system only the simpler achromat objectives are practical. With dark-ground illumination the range of objectives is considerably widened, and more elaborate lenses with higher apertures and better ‘light gathering power’ are possible.

Incident light fluorescence
The trend today in fluorescence techniques is in incident illumination, or lighting from above and through the objective down to the object ( Fig. 3.28 ). A number of advantages are gained over the transmitted route.

Figure 3.28 (a) Diagram of incident fluorescence microscope layout. (b) Effect of dichroic mirror.
In principle the excitation beam, after passing the selection filters, is diverted through the objective on to the preparation where fluorescence is stimulated. This fluorescence travels back to the observer by the normal route ( Fig. 3.28a ). Dichroic mirrors have been produced to divide and divert the beam. These mirrors have the property of being able to transmit light of some wavelengths and reflect other wavelengths ( Fig. 3.28b ). By selection of the appropriate mirror, the wavelength desired is reflected to the object; the remainder passes through to be lost. At the same time, visible fluorescent light collected by the objective in the normal way can pass to the eyepiece, and any excitation rays bouncing back (from slide and coverglass) are reflected back along their original path to the source, thus being prevented from reaching the observer. Since the objective in this system also acts as a condenser, the illumination and objective numerical apertures are one and the same, optically correct, and at their most efficient condition. Fluorescence is stimulated on the observer’s side of the preparation, and is therefore more brilliant, not being masked by covering material or section thickness.
It is also possible to bring into use any type of objective, including sophisticated phase contrast and interference contrast objectives, for simultaneous transmitted illumination with normal tungsten lighting, and to demonstrate both the fluorescence and the morphology of the preparation. This is useful where normal stains cannot be used for fear of masking any fluorescent reactions.
The use of dichroic mirrors in these systems has made possible much brighter images, since up to 90% of the exciting energy can reach the preparation, and 90% of the resultant visible light can be presented to the eye. In addition, new objectives, of both oil and water immersion types, in low and high powers, have been developed. As immersion objectives they have higher numerical apertures, and can gather more light, avoiding much of the lost stray light reflected from coverslips. The use of low-magnification eyepieces is now more widely accepted, improving fluorescence techniques far beyond anything hitherto possible.
Due to the filters and light sources used in fluorescence microscopy, modern systems generally rely on digital image capture, and these images are monochrome (black and white) images. The highly colored fluorescence images which appear in publications are all the result of pseudo-coloring composite images.

The confocal microscope
In fluorescence microscopy using conventional epi-fluorescence microscopes, the fluorochrome present in the field of view will be excited whether it is in or out of focus. The effect is that the out-of-focus fluorescence will reduce the contrast and resolution of the image. The confocal system uses a pinhole stop to observe the specimen in such a way as to exclude the out-of-focus portion of the image. In essence the axial resolution in the confocal system is greatly improved to 0.35 mm (in reflection) with additional small but important gains in lateral resolution, and therefore the method lends itself to optical sectioning. With modern computer technology and Windows ® -based software a series of optical sections can be recombined to create a 3D image of a cell or structure even with multiple labeling techniques.

Other techniques
A number of specialized techniques for use in fluorescence microscopy have appeared in recent years. One significant technique is fluorescence resonance energy transfer or FRET, in which the excitation of one fluorescence material is captured locally by a second material, causing fluorescence of the second material and quenching of the first. Many new techniques have been developed based on the availability of new fluorescence labels and dyes. Many of these new dyes have unique emission spectra, and may also be much more resistant to fading than earlier dyes. A promising new fluorescence label is quantum dots, a label composed on nanometer-sized particles of semiconductor metals. These quantum dots are excited over a wide range of wavelengths, and emit very narrow band light based on the size of the semiconductor particle. The advantage of quantum dots is that many different dots can be excited by the same excitation light, and each will emit a specific color based on the dot size.
The advent of powerful computers and sophisticated programs has led to many new approaches to microscopy. One specific technique that should be mentioned is multispectral imaging , in which a number of images are collected at narrow, specific wavelengths. Such images provide new information as individual components of the image can be easily enhanced, thus improving contrast or perceived resolution of specimen morphology.
In recent years, it has been demonstrated that images can be collected at resolutions that far exceed the resolution limits of traditional microscope lenses. These techniques require highly specialized microscopes equipped with exactingly precise scanning stages, both X and Y axis, as well as Z axis (focus) motorized functions, and in almost all cases require tiny pinholes in the optical system, similar to those used with confocal microscopy, although generally much smaller in size. These instruments are also used primarily to examine specimens that have been stained or treated with fluorescence compounds. One of the earliest demonstrations of this approach was that of Betzig et al. (1987) . This early demonstration proved the concept of Near Field Scanning Microscopy. More recently, a variety of ancillary approaches to super resolution have been developed. A summary of current approaches to super resolution was published by Huang (2010) .

Use of the microscope
The microscope itself should receive some attention before it is used. Is the illumination correctly centered? Is the condenser centered and in its proper position? Make sure that the objectives are firmly screwed home, and that the eyepieces are indeed a pair. If the microscope is set aside exclusively for your personal use, these things will probably be in order, but if the instrument is in communal use then it is likely that someone has altered or readjusted something to suit their own purpose. Above all, make sure that the optical parts are clean, and free from dust. Greasy fingerprints and dust are enemies of the optical glass.
Often when an immersion objective has been used, the next objective is swung into position, straight into the oil drop on the slide. Sometimes oil is left on the front of an immersion lens, forgotten for some time, and by the time it is used it has gathered some dust and formed a translucent film. Use oil only for an oil immersion objective. Keep it away from everything else. After a session with immersion objectives, clean them. Gross amounts of oil can be wiped off with lens tissue. The use of xylene for washing the front lens is now frowned upon. Petroleum spirit is recommended by some manufacturers. Alcohol and acetone should be avoided as they may seep into the mount and dissolve the cements.
The front lenses of some objectives are difficult to clean because of their concave shape, and many people recommend the use of swabs of cotton wool or tissue to remove the dirt from the concavity. A better method is the use of a piece of expanded polystyrene such as is used in the packing of delicate equipment and lamp bulbs. A freshly broken surface will be dust free and contain no hard crystals; it cannot scratch the glass. By pressing the polystyrene on to the lens and rotating it in the optical axis, grease, moisture, and dust are quickly removed, even from the deepest of concave lenses. Do not use polystyrene if the lens is wet with xylene, or it will coat the glass with a layer of dissolved plastic and make matters worse.
Eyepieces become coated with a fine film of grease from eyelashes. They should be cleaned from time to time with a lens tissue. Tissues are better than cloths as they are usually stored in small numbers in a protective packet, and are discarded after use. Cloths are liable to lie around on the bench for some time, picking up dust and other harmful substances.

Setting up the microscope

Centering the lamp
This is possible on most modern microscopes (some have pre-centered bulbs and do not require attention), either by a pair of centering screws acting against a spring, or by loosening a screw collar and orienting the lamp holder. A ground glass or plastic disc with concentric circles engraved upon it is placed in the light path, usually on the dust protection glass in the foot of the microscope. By adjusting the lamp condenser, or by moving the lamp in its mounting, the filament can be imaged on the ground-glass disc. It is then a simple matter to adjust the bulb position until the image is in the center.

Adjusting the condenser
If the microscope has an adjustable condenser, this is also a simple procedure. First, select a low-power objective (x10) and, with a suitable preparation on the stage, focus the specimen (to establish the plane of the object). Most condensers have a top lens that is normally swung into the light path for the higher-power objectives. This should be turned into position. Open the aperture diaphragm (in the condenser), close the field diaphragm (in the foot of the microscope) to a small aperture. An image of the field diaphragm should now be visible in the field of view. Adjust the height of the condenser until the image of the field diaphragm is sharply focused. This will be the correct position for slides of the same thickness. Now with the adjusting screws move the diaphragm image to the center. If the aperture is opened until it almost reaches the sides of the field of view, the centering is more accurate. When the correct position has been reached, open the field diaphragm until it just disappears from view. Adjust the aperture (or substage) iris diaphragm, by closing it down, removing an eyepiece and, while looking down the tube, open the diaphragm until it occupies one-third of the field of view. Replace the eyepiece. If the image is dark at the edge, open the aperture diaphragm a little more. This has now adjusted the numerical aperture of the condenser to approximately that of the objective in use and achieved the optimum resolution. The illumination should now be correctly adjusted and ready for use.

Inserting the slide
When changing slides it is always good practice to lower the stage before removing the slide. The risk of damaging the objective is reduced. When a new slide is placed on the stage and the objective lowered towards the focal plane, watch from the side and gently lower the objective (or, raise the stage) until it is almost in contact, then look into the eyepieces and complete the focusing by bringing the objective away from the preparation. If you look through the eyepiece as the objective approaches the slide, you stand a great chance of missing the focus plane and of crashing the objective through the slide with the possible destruction of both the objective and the specimen.
Make sure that the slide is the right way up. With the higher-power objectives, the working distance is very short and you may never find the image.

Additional resources
Many additional resources for microscopy may be found on the internet. A site containing much information as well as graphical examples is the Molecular Expressions Microscopy Primer: .


Betzig, et al., 1987. Collection mode near-field optical scanning microscopy. Appl Phys Lett 51, 2088–2090.
Huang, B., 2010. Super-resolution optical microscopy: multiple choices. Curr Opin Chem Biol (Feb), 14.
4 Fixation of tissues

Anthony Rhodes

Appropriate fixation of tissues for histological examination is extremely important. Without attention to this process, the range of tests performed in a modern histopathology laboratory will be rendered ineffective and practically useless. The concept of fixation of biological tissues in order to understand biological function and structure has led to the development of many types of fixatives over the last century. The mechanisms and principles by which specific fixatives act to harden and preserve tissues and prevent the loss of specific molecules fall into broad categories. These include the covalent addition of reactive groups and of cross-links, dehydration, and the effects of acids, salt formation, and heat, along with combinations of these actions. Compound fixatives may function via several of these mechanisms.
Although each fixative has advantages, they all have many disadvantages. These include molecular loss from ‘fixed’ tissues, swelling or shrinkage of tissues during the process, variations in the quality of histochemical and immunohistochemical staining, the ability to perform biochemical analysis accurately, and varying capabilities to maintain the structures of cellular organelles. One of the major problems with fixation using formaldehyde has been the loss of antigen immunorecognition due to that type of fixation combined with processing the tissue to paraffin wax ( Eltoum et al. 2001a , 2001b ). However, from a clinical perspective the advent of heat-induced epitope retrieval methods, instigated in the early 1990s, have overcome many of these limitations ( Shi et al. 1991 ). Similarly, the analysis of mRNA and DNA from formalin-fixed, paraffin-embedded tissue has been problematic ( Grizzle et al. 2001 ; Jewell et al. 2002 ; Steg et al. 2006 ; Lykidis et al. 2007 ). All widely used fixatives are selected by compromise; their good aspects are balanced against less desirable features. This chapter discusses the basics of fixation, the advantages and disadvantages of specific fixatives, and provides some of the formulae for specific fixatives currently used in pathology, histology, and anatomy.
The major objective of fixation in pathology is to maintain clear and consistent morphological features ( Eltoum et al. 2001a , 2001b ; Grizzle et al. 2001 ). The development of specific fixatives usually has been empirical, and much of the understanding of the mechanisms of fixation has been based upon information obtained from leather tanning and vaccine production. In order to visualize the microanatomy of a tissue, its stained sections must maintain the original microscopic relationships among cells, cellular components (e.g. the cytoplasm and nuclei), and extracellular material with little disruption of the organization of the tissue, and must maintain the tissue’s local chemical composition. Many tissue components are soluble in aqueous acid or other liquid environments, and a reliable view of the microanatomy and microenvironment of these tissues requires that the soluble components are not lost during fixation and tissue processing. Minimizing the loss of cellular components, including proteins, peptides, mRNA, DNA, and lipids, prevents the destruction of macromolecular structures such as cytoplasmic membranes, smooth endoplasmic reticulum, rough endoplasmic reticulum, nuclear membranes, lysosomes, and mitochondria. Each fixative, combined with the tissue processing protocol, maintains some molecular and macromolecular aspects of the tissue better than other fixative/processing combinations. For example, if soluble components are lost from the cytoplasm of cells, the color of the cytoplasm on hematoxylin and eosin (H&E) staining will be reduced or modified and aspects of the appearance of the microanatomy of the tissue, e.g. mitochondria, will be lost or damaged. Similarly, immunohistochemical evaluations of structure and function may be reduced or lost.
Almost any method of fixation induces shrinkage/swelling, hardening of tissues and color variations in various histochemical stains ( Sheehan & Hrapchak 1980 ; Horobin 1982 ; Fox et al. 1985 ; Carson 1990 ; Kiernan 1999 ; O’Leary & Mason 2004 ). Various methods of fixation always produce some artifacts in the appearance of tissue on staining; however, for diagnostic pathology it is important that such artifacts are consistent.
The fixative acts by minimizing the loss or enzymatic destruction of cellular and extracellular molecules, maintaining macromolecular structures and protecting tissues from destruction by microorganisms. This results in one view of a dynamically changing, viable tissue ( Grizzle et al. 2001 ). A fixative should also prevent the subsequent breakdown of the tissue or molecular features by enzymatic activity and/or by microorganisms during long-term storage, because diagnostic/therapeutic tissues removed from patients are important resources which may be re-analyzed in the future.
A fixative not only interacts initially with the tissue in its aqueous environment but also, subsequently, the unreacted fixative and the chemical modifications induced by the fixative continue to react. Fixation interacts with all phases of processing and staining from dehydration to staining of tissue sections using histochemical, enzymatic or immunohistochemical stains ( Eltoum et al. 2001b ; Rait et al. 2004 ). A stained tissue section produced after specific fixation combined with tissue processing produces a compromise in the picture that is formed of one or more features of the original living tissue. To date, a universal or ideal fixative has not been identified. Fixatives are therefore selected based on their ability to produce a final product needed to demonstrate a specific feature of a specific tissue ( Grizzle et al. 2001 ). In diagnostic pathology, the fixative of choice for most pathologists has been 10% neutral buffered formalin ( Grizzle et al. 2001 ).
The most important characteristic of a fixative is to support high quality and consistent staining with H&E, both initially and after storage of the paraffin blocks for at least a decade. The fixative must have the ability to prevent short- and long-term destruction of the micro-architecture of the tissue by stopping the activity of catabolic enzymes and hence autolysis, minimizing the diffusion of soluble molecules from their original locations. Another important characteristic of a good fixative, which helps maintain tissue and cellular integrity, is the inanimation of infectious agents, which helps maintain tissue and cellular integrity. It is also important to have good toxicological and flammability profiles that permit the safe use of the fixative ( Grizzle & Fredenburgh 2005 ). The advent of new biological methods, increased understanding of the human genome, and the need to rapidly evaluate the biology of disease processes, means that fixatives should also permit the recovery of macromolecules including proteins, mRNA, and DNA without extensive biochemical modifications from fixed and paraffin-embedded tissues.
Other important characteristics of an ideal fixative include being useful for a wide variety of tissues, including fatty, lymphoid, and neural tissues. It should preserve small and large specimens and support histochemical, immunohistochemical, in situ hybridization and other specialized procedures. It should penetrate and fix tissues rapidly, have a shelf life of at least one year, and be compatible with modern automated tissue processors. The fixative should be readily disposable or recyclable and support long-term tissue storage giving excellent microtomy of paraffin blocks, and should be cost- effective ( Dapson 1993 ).

Types of fixation
Fixation of tissues can be accomplished by physical and/or chemical methods. Physical methods such as heating, microwaving, and freeze-drying are independent processes and not used commonly in the routine practice of medical or veterinary pathology, anatomy, and histology, except for the use of dry heat fixation of microorganisms prior to Gram staining. Most methods of fixation used in processing of tissue for histopathological diagnoses rely on chemical fixation carried out by liquid fixatives. Reproducibility over time of the microscopic appearances of tissues after H&E staining is the prime requirement of the fixatives used for diagnostic pathology. Methods of fixation used in research protocols may be more varied, including fixation using vapors and fixation of whole animals by perfusing the animal’s vascular system with a fixative ( Eltoum et al. 2001a , 2001b ).
Several chemicals or their combinations can act as good fixatives, and accomplish many of the stated goals of fixation. Some fixatives add covalent reactive groups which may induce cross-links between proteins, individual protein moieties, within nucleic acids, and between nucleic acids and proteins ( Horobin 1982 ; Eltoum et al. 2001a , 2001b ; Rait et al. 2004 , 2005 ). The best examples of such ‘cross-linking fixatives’ are formaldehyde and glutaraldehyde. Another approach to fixation is the use of agents that remove free water from tissues and hence precipitate and coagulate proteins; examples of these dehydrants include ethanol, methanol, and acetone. These agents denature proteins by breaking the hydrophobic bonds which are responsible for the tertiary structure of proteins. Other fixatives, such as acetic acid, trichloroacetic acid, mercuric chloride, and zinc acetate, act by denaturing proteins and nucleic acids through changes in pH or via salt formation. Some fixatives are mixtures of reagents and are referred to as compound fixatives, e.g. alcoholic formalin acts to fix tissues by adding covalent hydroxymethyl groups and cross-links as well as by coagulation and dehydration.

Physical methods of fixation

Heat fixation
The simplest form of fixation is heat. Boiling or poaching an egg precipitates the proteins and, on cutting, the yolk and egg white can be identified separately. Each component is less soluble in water after heat fixation than the same component of a fresh egg. Picking up a frozen section on a warm microscope slide both attaches the section to the slide and partially fixes it by heat and dehydration. Even though adequate morphology could be obtained by boiling tissue in normal saline, in histopathology heat is primarily used to accelerate other forms of fixation as well as the steps of tissue processing.

Microwave fixation
Microwave heating speeds fixation and can reduce times for fixation of some gross specimens and histological sections from more than 12 hours to less than 20 minutes ( Anonymous 2001 ; Kok & Boon 2003 ; Leong 2005 ). Microwaving tissue in formalin results in the production of large amounts of dangerous vapors, so in the absence of a hood for fixation, or a microwave processing system designed to handle these vapors, this may cause safety problems. Recently, commercial glyoxal-based fixatives which do not form vapors when heated at 55°C have been introduced as an efficient method of microwave fixation.

Freeze-drying and freeze substitution
Freeze-drying is a useful technique for studying soluble materials and small molecules; tissues are cut into thin sections, immersed in liquid nitrogen, and the water is removed in a vacuum chamber at −40°C. The tissue can be post-fixed with formaldehyde vapor. In substitution, specimens are immersed in fixatives at −40°C, such as acetone or alcohol, which slowly remove water through dissolution of ice crystals, and the proteins are not denatured; bringing the temperature gradually to 4°C will complete the fixation process ( Pearse 1980 ). These methods of fixation are used primarily in the research environment and are rarely used in the clinical laboratory setting.

Chemical fixation
Chemical fixation utilizes organic or non-organic solutions to maintain adequate morphological preservation. Chemical fixatives can be considered as members of three major categories: coagulant, cross-linking, and compound fixatives ( Baker, 1958 ).

Coagulant fixatives
Both organic and non-organic solutions may coagulate proteins, making them insoluble. Cellular architecture is maintained primarily by lipoproteins and by fibrous proteins such as collagen; coagulating such proteins maintains tissue histomorphology at the light microscopic level. Unfortunately, because coagulant fixatives result in cytoplasmic flocculation as well as poor preservation of mitochondria and secretory granules, such fixatives are not useful in ultrastructural analysis.

Dehydrant coagulant fixatives
The most commonly used coagulating fixatives are alcohols (e.g. ethanol, methanol) and acetone. Methanol is closer to the structure of water than ethanol. Ethanol therefore competes more strongly than methanol in the interaction with hydrophobic areas of molecules; thus, coagulant fixation begins at a concentration of 50–60% for ethanol but requires a concentration of 80% or more for methanol ( Lillie & Fullmer 1976 ). Removal and replacement of free water from tissue by any of these agents has several potential effects on proteins within the tissue. Water molecules surround hydrophobic areas of proteins and, by repulsion, force hydrophobic chemical groups into closer contact with each other and hence stabilize hydrophobic bonding. By removing water, the opposite principle weakens hydrophobic bonding. Similarly, molecules of water participate in hydrogen bonding in hydrophilic areas of proteins; so removal of water destabilizes this hydrogen bonding. Together, these changes act to disrupt the tertiary structure of proteins. In addition, with the water removed, the structure of the protein may become partially reversed, with hydrophobic groups moving to the outside surface of the protein. Once the tertiary structure of a soluble protein has been modified, the rate of reversal to a more ordered soluble state is slow and most proteins after coagulation remain insoluble even if returned to an aqueous environment.
Disruption of the tertiary structure of proteins, i.e. denaturation, changes their physical properties, potentially causing insolubility and loss of function. Even though most proteins become less soluble in organic environments, up to 13% of protein may be lost, for example with acetone fixation ( Horobin 1982 ). Factors that influence the solubility of macromolecules include:

1. Temperature, pressure, and pH.
2. Ionic strength of the solute.
3. The salting-in constant, which expresses the contribution of the electrostatic interactions.
4. The salting-in and salting-out interactions.
5. The type(s) of denaturing reagent(s) ( Herskovits et al. 1970 ; Horobin 1982 ; Papanikolau & Kokkinidis 1997 ; Bhakuni 1998 ).
Alcohol denatures protein differently, depending on the choice and concentration of alcohol, the presence of organic and non-organic substances, and the pH and temperature of fixation. For example, ethanol denatures proteins > phenols > water and polyhydric alcohols > monocarboxylic acids > dicarboxylic acids ( Bhakuni 1998 ).

Other types of coagulant fixative
Acidic coagulants such as picric acid and trichloroacetic acid change the charges on the ionizable side chains, e.g. ( NH 2 → NH 3 + ) and (COO − → COOH), of proteins and disrupt electrostatic and hydrogen bonding. These acids also may insert a lipophilic anion into a hydrophilic region and hence disrupt the tertiary structures of proteins ( Horobin 1982 ). Acetic acid coagulates nucleic acids but does not fix or precipitate proteins; it is therefore added to other fixatives to prevent the loss of nucleic acids. Trichloroacetic acid (Cl 3 CCOOH) can penetrate hydrophobic domains of proteins and the anion produced ( C COO − ) reacts with charged amine groups. This interaction precipitates proteins and extracts nucleic acids. Picric acid or trinitrophenol slightly dissolves in water to form a weak acid solution (pH 2.0). In reactions, it forms salts with basic groups of proteins, causing the proteins to coagulate. If the solution is neutralized, precipitated protein may redissolve. Picric acid fixation produces brighter staining, but the low pH solutions of picric acid may cause hydrolysis and loss of nucleic acids

Non-coagulant cross-linking fixatives
Several chemicals were selected as fixatives secondary to their potential actions of forming cross-links within and between proteins and nucleic acids as well as between nucleic acids and proteins. Cross-linking may not be a major mechanism at current short times of fixation, and therefore ‘covalent additive fixatives’ may be a better name for this group. Examples include formaldehyde, glutaraldehyde, and other aldehydes, e.g. chloral hydrate and glyoxal, metal salts such as mercuric and zinc chloride, and other metallic compounds such as osmium tetroxide. Aldehyde groups are chemically and biologically reactive and are responsible for many histochemical reactions, e.g. free aldehyde groups may be responsible for argentaffin reactions ( Papanikolau & Kokkinidis 1997 ).

Formaldehyde fixation
Formaldehyde in its 10% neutral buffered form (NBF) is the most common fixative used in diagnostic pathology. Pure formaldehyde is a vapor that, when completely dissolved in water, forms a solution containing 37–40% formaldehyde; this aqueous solution is known as ‘formalin’. The usual ‘10% formalin’ used in fixation of tissues is a 10% solution of formalin; i.e., it contains about 4% weight to volume of formaldehyde. The reactions of formaldehyde with macromolecules are numerous and complex. Fraenkel-Conrat and his colleagues, using simple chemistry, meticulously identified most of the reactions of formaldehyde with amino acids and proteins ( French & Edsall 1945 ; Fraenkel-Conrat & Olcott 1948a , 1948b ; Fraenkel-Conrat & Mecham 1949 ). In an aqueous solution formaldehyde forms methylene hydrate, a methylene glycol as the first step in fixation ( Singer 1962 ).

Methylene hydrate reacts with several side chains of proteins to form reactive hydroxymethyl side groups ( CH 2 OH). If relatively short fixation times are used with 10% neutral buffered formalin (hours to days), the formation of hydroxymethyl side chains is probably the primary and characteristic reaction. The formation of actual cross-links may be relatively rare at the currently used relatively short times of fixation.
Formaldehyde also reacts with nuclear proteins and nucleic acids ( Kok & Boon 2003 ; Leong 2005 ). It penetrates between nucleic acids and proteins and stabilizes the nucleic acid-protein shell, and it also modifies nucleotides by reacting with free amino groups, as it does with proteins. In naked and free DNA, the cross-linking reactions are believed to start at adenine-thymidine (AT)-rich regions and cross-linking increases with increasing temperature ( McGhee & von Hippel 1975a , 1975b , 1977a , 1977b ). Formaldehyde reacts with C C and SH bonds in unsaturated lipids, but does not interact with carbohydrates ( French & Edsall 1945 ; Hayat 1981 ).
The side chains of peptides or proteins that are most reactive with methylene hydrate, and hence have the highest affinity for formaldehyde, include lysine, cysteine, histidine, arginine, tyrosine, and reactive hydroxyl groups of serine and threonine (see Table 4.1 ) ( Means & Feeney 1995 ).

Table 4.1 Action of major single or combination fixatives

Gustavson (1956) reported that one of the most important cross-links in ‘over-fixation’, i.e. in tanning, is that between lysine and the amide group of the protein backbone. Due to the shorter fixation times of current diagnostic pathological and biological applications, cross-linking reactions with the protein backbone are unlikely to occur ( French & Edsall 1945 ; Fraenkel-Conrat et al. 1945 , 1947 ; Fraenkel-Conrat & Olcott 1948a , 1948b ; Fraenkel-Conrat & Mecham 1949 ; Gustavson 1956 ).

Reversibility of formaldehyde-macromolecular reactions
The reactive groups may combine with hydrogen groups or with each other, forming methylene bridges. If the formalin is washed away, reactive groups may rapidly return to their original states, but any bridging that has already occurred may remain.
Washing for 24 hours removes about half of reactive groups, and 4 weeks of washing removes up to 90% ( Helander 1994 ). This suggests that actual cross-linking is a relatively slow process, so, in the rapid fixation used in diagnostic pathology, most ‘fixation’ with formaldehyde prior to tissue processing stops with the formation of reactive hydroxymethyl groups.
For long-term storage in formalin, the reactive groups may be oxidized to the more stable groups (e.g. acids NH COOH) which are not easily removed by washing in water or alcohol. Thus, following fixation, returning the specimen to water or alcohol further reduces the fixation of the specimen, because the reactive groups produced by the initial reaction with formalin may reverse and be removed. Although it was initially thought that cross-linking was most important in the fixation of tissue for biological uses (based on the limited number of cross-links over short periods of fixation), it is likely that formation of these hydroxymethyl groups actually denatures macromolecules and renders them insoluble. As these washing experiments have not been reproduced, the actual mechanisms and their importance to fixation by formaldehyde are uncertain. As well as simple washing under running water, over-fixation of tissue may be partially corrected by soaking the tissue in concentrated ammonia plus 20% chloral hydrate ( Lhotka & Ferreira 1949 ). Fraenkel-Conrat and his colleagues frequently noted that the addition and condensation reactions of formaldehyde with amino acids and proteins were unstable and could be reversed easily by dilution or dialysis ( Fraenkel-Conrat et al. 1945 , 1947 ; Fraenkel-Conrat & Olcott 1948a , 1948b ; Fraenkel-Conrat & Mecham 1949 ).
The principal type of cross-link in short-term fixation is thought to be between the hydroxymethyl group on a lysine side chain and arginine (through secondary amino groups), asparagine, glutamine (through secondary amide groups), or tyrosine (through hydroxyl group) ( Tome et al. 1990 ). For example, a lysine methyl hydroxyl amine group can react with an arginine group to form a lysine CH 2 –arginine cross-link; similarly, a tyrosine methyl hydroxyl amine group can bind with a cysteine group to form a tyrosine CH 2 –cysteine cross-link. Each of these cross-links between macromolecules has a different degree of stability, which can be modified by the temperature, pH, and type of the environment surrounding and permeating the tissue ( Eltoum et al. 2001b ). The time to saturation of human and animal tissues with active groups by formalin is about 24 hours, but cross-linking may continue for many weeks ( Helander 1994 ).
When formaldehyde dissolves in an unbuffered aqueous solution, it forms an acid solution (pH 5.0–5.5) because 5–10% of commercially available formaldehyde is formic acid. Acid formalin may react more slowly with proteins than NBF because amine groups become charged (e.g. N + H 3 ). In solution, this requires a much lower pH than 5.5. However, the requirement for a lower pH to produce N + H 3 groups may not be equivalent to that required in peptides. Acid formalin also preserves immunorecognition much better than NBF ( Arnold et al. 1996 ), and indeed the success of Taylor in the early days of immunocytochemistry to demonstrate immunoglobulins in paraffin-processed tissue sections, most probably relied on the fixation of the tissues in acid formalin ( Taylor et al. 1974 ). The disadvantage of using acid formalin for fixation is the formation of a brown-black pigment with degraded hemoglobulin. This heme-related pigment, which forms in tissue, is usually not a great problem unless patients have a blood abnormality (e.g. sickle cell disease, malaria).
Formaldehyde primarily preserves peptide-protein bonds and the general structure of cellular organelles. It can interact with nucleic acids but has little effect on carbohydrates and preserves lipids if the solutions contain calcium ( Bayliss High & Lake 1996 ).

Glutaraldehyde fixation
Less is known about glutaraldehyde’s biological reactions and effects compared to formaldehyde, as it has not been used as widely in biological applications. Glutaraldehyde is a bifunctional aldehyde that probably combines with the same reactive groups as does formaldehyde. In aqueous solutions glutaraldehyde polymerizes, forming cyclic and oligomeric compounds ( Hopwood 1985 ), and it is also oxidized to glutaric acid. To aid in stability, it requires storage at 4°C and at a pH of around 5 ( Hopwood 1969 ).
Unlike formaldehyde, glutaraldehyde has an aldehyde group on both ends of the molecule. With each reaction of the first group, an unreacted aldehyde group may be introduced into the protein and these aldehyde groups can act to further cross-link the protein. Alternatively, the aldehyde groups may react with a wide range of other histochemical targets, including antibodies, enzymes, or proteins. The reaction of glutaraldehyde with an isolated protein, such as bovine serum albumin, is fastest at pH 6–7, and is faster ( Habeeb 1966 ), and results in more cross-linking than formaldehyde ( Habeeb 1966 ; Hopwood 1969 ). Cross-linking is irreversible and withstands acids, urea, semicarbazide, and heat ( Hayat 1981 ). Like formaldehyde, reactions with lysine are the most important for forming cross-links.
Extensive cross-linking by glutaraldehyde results in better preservation of ultrastructure, but this method of fixation negatively affects immunohistochemical methods and slows the penetration by the fixative. Thus, any tissue fixed in glutaraldehyde must be small (0.5 mm maximum) and, unless the aldehyde groups are blocked, increased background staining will result if several histochemical methods are used ( Grizzle 1996a ). Glutaraldehyde does not react with carbohydrates or lipids unless they contain free amino groups as are found in some phospholipids ( Hayat 1981 ). At room temperature glutaraldehyde does not cross-link nucleic acids in the absence of nucleohistones but it may react with nucleic acids at or above 45°C ( Hayat 1981 ).

Osmium tetroxide fixation
Osmium tetroxide (OsO 4 ), a toxic solid, is soluble in water as well as non-polar solvents and can react with hydrophilic and hydrophobic sites including the side chains of proteins, potentially causing cross-linking ( Hopwood et al. 1990 ). The reactive sites include sulfydryl, disulfide, phenolic, hydroxyl, carboxyl, amide, and heterocyclic groups. Osmium tetroxide is known to interact with nucleic acids, specifically with the 2,3-glycol moiety in terminal ribose groups and the 5,6 double bonds of thymine residues. Nuclei fixed in OsO 4 and dehydrated with alcohol may show prominent clumping of DNA. This unacceptable artifact can be prevented by pre-fixation with potassium permanganate (KMnO 4 ), post-fixation with uranyl acetate, or by adding calcium ions and tryptophan during fixation ( Hayat 1981 ). The reaction of OsO 4 with carbohydrates is uncertain ( Hayat 1981 ). Large proportions of proteins and carbohydrates are lost from tissues during osmium fixation; some of this may be due to the superficial limited penetration of OsO 4 (i.e. <1 mm) into tissues or its slow rates of reaction. In electron microscopy, this loss is minimized by initial fixation of tissue in glutaraldehyde.
The best characterized reaction of osmium is its reaction with unsaturated bonds within lipids and phospholipids. In this reaction, osmium in its +8 valence state converts to a +6 valence state, which is colorless. If two unsaturated bonds are close together there may be cross-linking by OsO 4 . Although the complex is colorless at this point, the typical black staining of membranes expected from fixation with osmium requires the production of osmium dioxide (OsO 2 ·2H 2 O). Osmium dioxide is black, electron dense, and insoluble in aqueous solution; it precipitates as the above unstable compounds break down and becomes deposited on cellular membranes. The breakdown of osmium +6 valence complexes to osmium dioxide (+4 valence state) is facilitated by a reaction with solutions of ethanol.
In addition to its use as a secondary fixative for electron microscope examinations, OsO 4 can also be used to stain lipids in frozen sections. Osmium tetroxide fixation causes tissue swelling which is reversed during dehydration steps. Swelling can also be minimized by adding calcium or sodium chloride to osmium-containing fixatives ( Hayat 1981 ).

Cross-linking fixatives for electron microscopy
Cell organelles such as cytoplasmic and nuclear membranes, mitochondria, membrane-bound secretory granules, and smooth and rough endoplasmic reticulum need to be preserved carefully for electron microscopy. The lipids in these structures are extracted by many fixatives with dehydrants (e.g. alcohols). Therefore for ultrastructural examination it is important to use a fixative that does not solubilize lipids. The preferred fixatives are a strong cross-linking fixative such as glutaraldehyde, a combination of glutaraldehyde and formaldehyde, or Carson’s modified Millonig’s, followed by post-fixation in an agent that further stabilizes as well as emphasizes membranes such as OsO 4 .

Mercuric chloride
Historically, mercuric chloride was greatly favored for its qualities of enhancing the staining properties of tissues, particularly for trichrome stains. However, it is now rarely used in the clinical laboratory due to the health and safety issues involved with the use of a mercury-containing fixative, and also due to the reduced reliance on ‘special stains’. A further major disadvantage of mercuric chloride fixation is the inevitable formation of deposits of intensely black precipitates of mercuric pigment in the tissues. This subsequently gives them inferior value for immunohistochemical and molecular studies. In recently fixed tissues, these precipitates can be readily removed by a Lugol’s iodine step in the staining procedure, followed by bleaching of the section in sodium hypochlorite solution (Hypo). However, this is not effective on mercuric chloride fixed tissues which have been stored for a number of years as paraffin blocks. In these tissues, retrospective analysis by immunohistochemistry and molecular techniques becomes unreliable due to the formation of much larger aggregates of mercuric pigment which cannot be removed subsequently by Lugol’s iodine. The chemistry of fixation using mercuric chloride is not well understood. It is, however, known that mercuric chloride reacts with ammonium salts, amines, amides, amino acids, and sulfydryl groups, and hardens tissues. It is especially reactive with cysteine, forming a dimercaptide ( Hopwood 2002 ) and acidifying the solution:

If only one cysteine is present, a reactive group of R S Hg Cl is likely.
Mercury-based fixatives are toxic and should be handled with care. They should not be allowed to come into contact with metal, and should be dissolved in distilled water to prevent the precipitation of mercury salts. Mercury-containing chemicals are an environmental disposal problem. These fixatives penetrate slowly, so specimens must be thin, and mercury and acid formaldehyde hematein pigments may deposit in tissue after fixation. Mercury fixatives ( Hopwood 1973 ) are no longer used routinely except by some laboratories for fixing hematopoietic tissues (especially B5). A potential replacement for mercuric chloride is zinc sulfate. Special formulations of zinc sulfate in formaldehyde replacing mercuric chloride in B5 may give better nuclear detail than formaldehyde alone and improve tissue penetration ( Carson 1990 ).

Special fixatives

Dichromate and chromic acid fixation
Chromium trioxide dissolves in water to produce an acidic solution of chromic acid, with a pH of 0.85. Chromic acid is a powerful oxidizing agent which produces aldehyde from the 1, 2-diglycol residues of polysaccharides. These aldehydes can react in histochemical stains (PAS and argentaffin/argyrophil) and should increase the background of immunohistochemical staining ( Grizzle 1996a ).
Actual chromic salts (i.e. chromium ions in +3 valence state) may destroy animal tissues ( Kiernan 1999 ) but chromium ions in their +6 state coagulate proteins and nucleic acids. The fixation and hardening reactions are not understood completely but probably involve the oxidation of proteins, which varies in strength depending upon the pH of the fixative, plus interaction of the reduced chromate ions directly in cross-linking proteins ( Pearse & Stoward 1980 ). Chromium ions specifically interact with the carboxyl and hydroxyl side chains of proteins. Chromic acid also interacts with disulfide bridges and attacks lipophilic residues such as tyrosine and methionine ( Horobin 1982 ). Fixatives containing chromate at a pH of 3.5–5.0 make proteins insoluble without coagulation. Chromate is reported to make unsaturated but not saturated lipids insoluble upon prolonged (>48 hours) fixation and hence mitochondria are well preserved by dichromate fixatives.
Dichromate-containing fixatives have primarily been used to prepare neuroendocrine tissues for staining, especially normal adrenal medulla and related tumors (e.g. phaeochromocytomas). However, reliance on the chromaffin reaction used to identify chromaffin granules following dichromate fixation has greatly diminished, being replaced by immunohistochemistry to a range of neuroendocrine markers, to include neuron-specific enolase, chromagranin A, and synaptophysin ( Grizzle 1996a , 1996b ).

Fixatives for DNA, RNA, and protein analysis
Lykidis et al. (2007) conducted a comprehensive analysis of 25 fixative compounds, many reputed to provide improved preservation of DNA and RNA and proteins in tissues for immunocytochemical analysis, whilst at the same time ensuring optimal morphological preservation. The compounds included the commercially available HOPE (HEPES-glutamic acid buffer mediated Organic Solvent Protection Effect) fixative and the reversible cross-linker dithio-bis[succinimidyl propionate] (DSP) for immunocytochemistry and expression profiling, in addition to zinc-based fixatives. They concluded that a novel zinc formation (Z7) containing zinc trifluoroacetate, zinc chloride and calcium acetate was significantly better than the standard zinc-based fixative (Z2) and NBF for DNA, RNA and antigen perseveration. DNA and RNA fragments up to 2.4kb and 361bp in length, respectively, were detected by PCR, reverse transcriptase PCR and real-time PCR in the Z7 fixed tissues, in addition to allowing for protein analysis using 2D electrophoresis. Nucleic acids and protein were found to be stable over a period of 6–14 months. Moreover, the fixative is less toxic than formaldehyde formulations. Whilst this fixative appears to show great promise, it should be borne in mind that fixation in NBF will also allow the extraction of similarly sized fragments of DNA and RNA for analysis by PCR-based technologies, within this time frame.

Metallic ions as a fixative supplement
Several metallic ions have been used as aids in fixation, including Hg 2+ , Pb 2+ , Co 2+ , Cu 2+ , Cd 2+ , [UO 2 ] 2+ , [PtCl 6 ] 2+ , and Zn 2+ . Mercury, lead, and zinc are used most commonly in current fixatives, e.g. zinc-containing formaldehyde is suggested to be a better fixative for immunohistochemistry than formaldehyde alone. This does however depend upon the pH of the formaldehyde, as well as the zinc formaldehyde ( Arnold et al. 1996 ; Eltoum et al. 2001a ).

Compound fixatives
Pathologists use formaldehyde-based fixatives to ensure reproducible histomorphometric patterns. Other agents may be added to formaldehyde to produce specific effects that are not possible with formaldehyde alone. The dehydrant ethanol, for example, can be added to formaldehyde to produce alcoholic formalin. This combination preserves molecules such as glycogen and results in less shrinkage and hardening than pure dehydrants.
Compound fixatives are useful for specific tissues, e.g. alcoholic formalin for fixation of some fatty tissues, such as breast, in which preservation of the lipid is not important. In addition, fixation of gross specimens in alcoholic formalin may aid in identifying lymph nodes embedded in fat. Some combined fixatives including alcoholic formalin are good at preserving antigen immunorecognition, but non-specific staining or background staining in immunohistochemical procedures can be increased. Unreacted aldehyde groups in glutaraldehyde-formaldehyde fixation for example may increase background staining, and alcoholic formalin may cause non-specific staining of myelinated nerves ( Grizzle et al. 1995 , 1997 , 1998a , 1998b ; Arnold et al. 1996 ; Grizzle 1996b ).

Factors affecting the quality of fixation

Buffers and pH
The effect of pH on fixation with formaldehyde may be profound depending upon the applications to which the tissues will be exposed. In a strongly acidic environment, the primary amine target groups ( NH 2 ) attract hydrogen ions ( NH + 3 ) and become unreactive to the hydrated formaldehyde (methylene hydrate or methylene glycol), and carboxyl groups ( COO − ) lose their charges ( COOH). This may affect the structure of proteins. Similarly the hydroxyl groups of alcohols ( OH) including serine and threonine may become less reactive in a strongly acidic environment. The extent of formation of reactive hydroxymethyl groups and cross-linking is reduced in unbuffered 4% formaldehyde ( Means & Feeney 1995 ), which is slightly acidic ( French & Edsall 1945 ), because the major methylene cross-links are between lysine and the free amino group on side chains. The decrease in the effectiveness of formaldehyde fixation and hence cross-linking in such a slightly acid environment has led some authors to suggest that unbuffered formalin is a better fixative than NBF with respect to immunorecognition of many antigens ( Arnold et al. 1996 ; Eltoum et al. 2001b ). This no doubt aided the detection of antigens before the early 1990s, prior to the advent of heat-induced epitope retrieval methods in immunocytochemistry. However, minimal delay in effectively fixing very labile antigens, such as the estrogen receptor, is vital in the immunohistochemical testing for a range of clinically important prognostic and predictive biomarkers. Whilst formaldehyde fixation remains the recommended method for optimal preservation of morphological features, proteins and nucleic acids in a clinical environment, the most reliable way of achieving optimal formalin fixation is through its buffering at pH 7.2–7.4 (i.e. neutral buffered formalin).
At the acidic pH of unbuffered formaldehyde, hemoglobin metabolic products are chemically modified to form a brown-black, insoluble, crystalline, birefringent pigment. The pigment forms at a pH of less than 5.7, and the extent of its formation increases in the pH range of 3.0 to 5.0. Formalin pigment is recognized easily and should not affect diagnoses except in patients with large amounts of hemoglobin breakdown products secondary to hematopoietic diseases. The pigment is removed easily with an alcoholic solution of picric acid. To avoid the formation of formalin pigment, neutral buffered formalin is used as the preferred formaldehyde-based fixative.
Acetic acids and other acids work mainly through lowering pH and disrupting the tertiary structure of proteins. Buffers are used to maintain optimum pH. The choice of specific buffer depends on the type of fixative and analyte. Commonly used buffers are phosphate, cacodylate, bicarbonate, Tris, and acetate. It is necessary to use low salt-buffered formalin in the new complex tissue processors in order to keep the machine ‘clean’, and reduce problems in its operation.

Duration of fixation and size of specimens
The factors that govern diffusion of a fixative into tissue were investigated by Medawar (1941) . He found that the depth (d) reached by a fixative is directly proportional to the square root of duration of fixation (t) and expressed this relation as:

The constant (k) is the coefficient of diffusability, which is specific to each fixative. Examples are 0.79 for 10% formaldehyde, 1.0 for 100% ethanol, and 1.33 for 3% potassium dichromate ( Hopwood 1969 ). Thus, for most fixatives, the time of fixation is approximately equal to the square of the distance which the fixative must penetrate. Most fixatives, such as NBF, will penetrate tissue to the depth of approximately 1 mm in one hour; hence for a 10 mm sphere, the fixative will not penetrate to the center until (5) 2 or 25 hours of fixation. It is important to note that the components of a compound fixative will penetrate the tissue at different rates, so that these aspects of the fixative will be best manifest in thin specimens.
Gross specimens should not rest on the bottom of a container of fixative: they should be separated from the bottom by wadded fixative-soaked paper or cloth, so allowing penetration of fixative or processing fluids from all directions. In addition, unfixed gross specimens which are to be cut and stored in fixative prior to processing should not be thicker than 0.5cm. When surgical specimens are to be processed to paraffin blocks, the time of penetration by fixative is more critical. Specific issues related to the processing of tissues have been reviewed by Grizzle et al. (2001) and Jones et al. (2001 ).
Fixation proceeds slowly and the period between the formation of reactive hydroxymethyl groups and the formation of a significant number of cross-links is unknown. Ninety percent of reactive groups can be removed by 4 weeks of washing ( Helander 1994 ), confirming that cross-linking is not a rapid process and may require weeks for completion of potential bonds.
Proteins inactivate fixatives, especially those in blood or bloody fluids. Bloody gross specimens should therefore be washed with saline prior to being put into fixative. The fixative volume should be at least 10 times the volume of the tissue specimen for optimal, rapid fixation. Currently in some laboratories, thin specimens may be fixed in NBF for only 5–6 hours including the short time of fixation in tissue processors. The extent of formation of cross-links during such rapid NBF ‘fixation’ is uncertain. Consequently, the formation of hydroxymethyl groups may predominate, as opposed to more resilient cross-linking. It has been suggested that rapid fixation is acceptable as long as histochemical staining remains adequate; and that immunohistochemistry and other molecular techniques are probably enhanced by shorter times of fixation using an aldehyde (e.g. formaldehyde)-based fixation. However, recent studies investigating the time taken to adequately fix clinical cases of breast cancer tissue for subsequent immunohistochemical detection of estrogen receptors illustrate that this practice can be detrimental to the optimal preservation of important antigens and should be avoided. Goldstein et al. (2003) found that 6–8 hours was the minimum time required to adequately fix breast tissue for immunohistochemical testing of estrogen receptors, regardless of the size and type of specimen. Consequently, the current guidelines for estrogen receptor and progesterone receptor testing produced by the American Society of Clinical Oncology (ASCO) and College of American Pathologists (CAP) recommend this minimal fixation time in neutral buffered formalin for all clinical breast cancer specimens ( Hammond et al. 2010 ).

Temperature of fixation
The diffusion of molecules increases with rising temperature due to their more rapid movement and vibration; i.e. the rate of penetration of a tissue by formaldehyde is faster at higher temperatures. Microwaves therefore have been used to speed formaldehyde fixation by both increasing the temperature and molecular movements. Increased vapor levels, however, are a safety problem ( Grizzle & Fredenburgh 2001 , 2005 ). Most chemical reactions occur more rapidly at higher temperatures and therefore formaldehyde reacts more rapidly with proteins ( Hopwood 1985 ). Closed tissue processors have their processing retort directly above the paraffin holding stations which are held at 60–65°C, making the retort slightly warmer than room temperature.

Concentration of fixative
Effectiveness and solubility primarily determine the appropriate concentration of fixatives. Concentrations of formalin above 10% tend to cause increased hardening and shrinkage ( Fox et al. 1985 ). In addition, higher concentrations result in formalin being present in its polymeric form, which can be deposited as white precipitate, as opposed to its monomeric form HO(H 2 CO)H, which at 4% provides for greatest solubility ( Baker 1958 ). Ethanol concentrations below 70% do not remove free water from tissues efficiently.

Osmolality of fixatives and ionic composition
The osmolality of the buffer and fixative is important; hypertonic and hypotonic solutions lead to shrinkage and swelling, respectively. The best morphological results are obtained with solutions that are slightly hypertonic (400–450 mOsm), though the osmolality for 10% NBF is about 1500 mOsm. Similarly, various ions (Na + , K + , Ca 2+ , Mg 2+ ) can affect cell shape and structure regardless of the osmotic effect. The ionic composition of fluids should be as isotonic as possible to the tissues.

The addition of electrolytes and non-electrolytes to fixatives improves the morphology of the fixed tissue. These additives include calcium chloride, potassium thiocyanate, ammonium sulfate, and potassium dihydrogen phosphate. The electrolytes may react either directly with proteins causing denaturation, or independently with the fixatives and cellular constituents ( Hayat 1981 ). The choice of electrolytes to be added to fixatives used on a tissue processor may vary. Fixatives buffered with electrolytes such as phosphates may cause problems with some processors due to precipitation of the salts. The addition of non-electrolyte substances such as sucrose, dextran, and detergent has also been reported to improve fixation ( Hayat 1981 ).

Selecting or avoiding specific fixatives
The choice of a fixative is a compromise, balancing their beneficial and detrimental effects. Kiernan (1999) originally produced a table of the actions of fixatives; this was later modified and published by Eltoum et al. (2001b) , and Table 4.1 is a further modification of the latter.
However, specific fixatives are unsuitable for most uses and should be avoided. The main problem with fixatives used in histological staining is the loss by solution/extraction of molecules that are targets of specific histochemical methods. Typically, some molecules are soluble in aqueous fixatives (e.g. glycogen), while others are soluble in organic-based fixatives (e.g. lipids). Some fixatives may chemically modify targets of histochemical staining and thus affect the quality of special stains (e.g. glutaraldehyde for silver stains); this includes modification of staining secondary to changes in pH induced by fixation. A good discussion of the effects of fixation on histochemistry is by Sheehan and Hrapchak (1980) .
The table of Sheehan and Hrapchak (1980) modified by ( Eltoum et al. 2001b ) has been changed so that harmful methods of fixation could be identified rapidly. Table 4.2 of this chapter is a further modification of the table.

Table 4.2 Incompatible stains and fixatives

Fixation for selected individual tissues

The globe must be firmly fixed in order to cut good sections for embedding. Eyes may be fixed in NBF, usually for about 48 hours; to speed fixation one or two small windows can be cut into the globe (avoid the retina and iris) after 24 hours. After gross description, the anterior (iris) and posterior (e.g. optic nerve) are removed with a new, sharp razor blade and the components of the globe are fixed for an additional 48 hours, or more, in buffered formaldehyde, before being processed. Embedding may be in celloidin or paraffin. Perfusion fixation of the eye is recommended for studies of the canal of Schlemm and/or the aqueous outflow pathways.

The problem of fixing a whole brain is to render it firm enough to investigate the neuroanatomy and to produce sections to show histopathology and to respond to immunochemistry if required. Conventionally this fixation takes at least 2 weeks. Adickes et al. (1997) proposed a perfusion technique which allows all of the above to be accomplished and the report issued in 5–6 days. This method depends on the perfusion of the brain via the middle cerebral arteries. Fixatives may also be enhanced by the use of microwave technology ( Anonymous 2001 ; Kok & Boon 2003 ; Leong 2005 ). Alcoholic formalin should not be used for fixation if immunohistochemistry is to be performed using biotin-avidin (streptavidin) methods (Grizzle et al., unpublished data).

Clinical samples should be fixed in 10% NBF for between a minimum of 6–8 hours and a maximum of 72 hours, and should be sliced at 5mm intervals after appropriate gross inspection and margins designation. Time from tissue acquisition to fixation should be as short as possible in order to prevent lysis of clinically important biomarkers, such as estrogen receptors, progesterone receptors and the human epidermal growth factor receptor-2 (HER2). They should be placed in a sufficient volume of NBF to allow adequate tissue penetration. If the tumor specimen has come from a remote geographical location, it should be bisected through the tumor on removal and sent to the laboratory immersed in a sufficient volume of NBF ( Hammond et al., 2010 ).

Lung biopsies are typically fixed in NBF. The lungs from autopsies may be inflated by and fixed in NBF via the trachea or major bronchi, and in our experience these lungs can be cut within 2 hours. Gross sections are fixed overnight and sections to be processed and cut the next day.

Lymphoid tissue
Special care should be taken with all lymphoid tissue, as many organisms (e.g. Mycobacterium tuberculosis and viruses) may sequester themselves in the lymphoid reticular system. The lymphoid tissue is usually sliced and a representative sample of fresh tissue taken for special studies (e.g. flow cytometry or molecular analysis). The rest of the lymph node is fixed in NBF, though some laboratories fix part of the tissue in B5 or zinc.

Biopsies of the testes are fixed routinely in NBF.

Muscle biopsies
Biopsies of muscle are received fresh. A portion is separated for enzyme histochemistry. The tissue for routine histological assessment is fixed in NBF and embedded so the fibers of the specimens are viewed in cross-section and longitudinally. After processing this is stained with H&E, a trichrome stain, and Congo red if amyloid is suspected.

Renal biopsies
Renal core biopsies should be subdivided into three and each piece should contain adequate numbers of glomeruli. Each portion is then preserved, depending upon the method to be used for analysis:

• NBF for routine histology
• Buffered glutaraldehyde (pH 7.3) for ultrastructural analysis
• Snap frozen in isopentane and liquid nitrogen for immunofluorescence examination.

Useful formulae for fixatives
Gray (1954) lists over 600 formulations for various fixatives. The following is a list of the fixatives and formulae most commonly used by biomedical scientists and histotechnologists. Many of these formulae are based on those presented in standard textbooks of histochemistry ( Sheehan & Hrapchak 1980 ; Carson 1990 ; Kiernan 1999 ). They vary slightly from text to text, but these variations are unlikely to cause problems.
For routine histology, 10% neutral buffered formalin (NBF) is frequently used for initial fixation and for the first station on tissue processors. NBF is composed of a 10% solution of phosphate buffered formaldehyde. Formaldehyde is commercially supplied as a 37–40% solution and in the following formulae is referred to as 37% formaldehyde.

Neutral buffered 10% formalin

Tap water 900 ml Formalin (37% formaldehyde solution) 100 ml Sodium phosphate, monobasic, monohydrate 4 g Sodium phosphate, dibasic, anhydrous 6.5 g
The pH should be 7.2–7.4
There are other formulations of NBF and related fixatives. NBF purchased from commercial companies may vary widely in its aldehyde content, and commercial companies may add material such as methanol ( Fox et al. 1985 ) or other agents to stabilize NBF preparations.

Carson’s modified Millonig’s phosphate buffered formalin

Formaldehyde (37–40%) 10 ml Tap water 90 ml Sodium phosphate, monobasic 1.86 g Sodium hydroxide 0.42 g
Deionized water can be used if tap water is hard and/or contains solids. The pH should be 7.2–7.4. This formula is reported to be better for ultrastructural preservation than NBF.
Sometimes the term ‘formal’ is used to refer to 10% formalin or 37% formaldehyde.

Formal (10% formalin), calcium acetate

Tap water 900 ml Formaldehyde (37%) 100 ml Calcium acetate 20 g
This is a good fixative for preservation of lipids.

Formal (10% formalin), saline

Tap water 900 ml Formaldehyde (37%) 100 ml Sodium chloride 9 g

Formal (10% formalin), zinc, unbuffered

Tap water 900 ml Formaldehyde (37%) 100 ml Sodium chloride 4.5 g Zinc chloride or (zinc sulfate) 1.6 g (or 3.6 g)
Zinc formalin is reported to be an excellent fixative for immunohistochemistry.

Formalin, buffered saline

Tap water 900 ml Formaldehyde (37%) 100 ml Sodium chloride 9 g Sodium phosphate, dibasic 12 g

Formalin, buffered zinc

10% neutral buffered formalin 1000 ml Zinc chloride 1.6 g

Mercuric fixatives
A problem with fixation in mercury solutions is that several types of pigment may combine with the mercury. These pigments are removed from sections by using iodine treatment followed by sodium thiosulfate.

Zenker’s solution

Distilled water 250 ml Mercuric chloride 12.5 g Potassium dichromate 6.3 g Sodium sulfate 2.5 g
Just before use add 5 ml of glacial acetic acid to 95 ml of above solution. This is a good fixative for bloody (congested) specimens and trichrome stains.

Helly’s solution

Distilled water 250 ml Mercuric chloride 12.5 g Potassium dichromate 6.3 g Sodium sulfate 2.5 g
Just before use add 5 ml of 37% formaldehyde to 95 ml of above solution. It is excellent for bone marrow extramedullary hematopoiesis and intercalated discs.

Schaudinn’s solution

Distilled water 50 ml Mercuric chloride 3.5 g Absolute ethanol 25 ml

Ohlmacher’s solution

Absolute ethanol 32 ml Chloroform 6 ml Glacial acetic acid 2 ml Mercuric chloride 8 g
This fixative penetrates rapidly.

Carnoy-Lebrun solution

Absolute ethanol 15 ml Chloroform 15 ml Glacial acetic acid 15 ml Mercuric chloride 8 g
This fixative penetrates rapidly.

B5 fixative

Stock solution:   Mercuric chloride 12 g Sodium acetate 2.5 g Distilled water 200 ml
Add 2 ml of formaldehyde (37%) to 20 ml of stock solution just before use.
Frequently used for bone marrow, lymph nodes, spleen, and other hematopoietic tissues.

Dichromate fixatives
There is a variation among the names attributed to the formulae of dichromate fixatives but not in the formulae themselves. Time of fixation (24 hours) is critical for dichromate fixatives. Tissue should be washed after fixation and transferred to 70% ethanol. Failure to wash the tissue after fixation may cause pigments to be precipitated. Extensive shrinkage occurs when tissues are processed to paraffin blocks.

Miller’s or Möller’s solution

Potassium dichromate 2.5 g Sodium sulfate 1 g Distilled water 100 ml

Möller’s or Regaud’s solution

Potassium dichromate 3 g Distilled water 80 ml
At time of use add 20 ml of formaldehyde (37%).

Orth’s solution

Potassium dichromate 2.5 g Sodium sulfate 1 g Distilled water 100 ml
At time of use add 10 ml of formaldehyde (37%).

Lead fixatives
See special fixatives .

Picric acid fixatives
Many picric acid fixatives require a saturated aqueous solution of picric acid. Aqueous picric acid 2.1% will produce a saturated solution and 5% picric acid a saturated solution in absolute ethanol.

Bouin’s solution

Saturated aqueous solution of picric acid 1500 ml Formaldehyde (37%) 500 ml Glacial acetic acid 100 ml
Bouin’s solution is an excellent general fixative for connective tissue stains. The yellow color can be removed with 70% ethanol, lithium carbonate, or another acid dye, separately or during the staining sequence. Bouin’s solution destroys membranes; therefore intact nuclei cannot be recovered from Bouin’s fixed tissue and there may be extensive shrinkage of larger specimens.

Hollande’s solution

Distilled water 1000 ml Formaldehyde (37%) 100 ml Acetic acid 15 ml Picric acid 40 g Copper acetate 25 g
A useful fixative for gastrointestinal biopsies and endocrine tissue; specimens are washed before exposure to NBF.

Dehydrant fixatives
Dehydrant fixatives act to remove free and bound water, causing a change to the tertiary structure of proteins so that they precipitate, leaving the nucleic acids relatively unchanged. Ultrastructure is destroyed by any of these four dehydrants due to the extraction of lipids, and each may cause excessive shrinking of tissue components after more than 3–4 hours of fixation. Each of these fixatives can be modified by adding other chemicals to produce specific effects.

1. Ethanol, absolute
2. Ethanol, 95%
3. Ethanol, 70–95%
4. Methanol, 100%
5. Acetone, 100%
Methanol is useful for touch preparations and smears, especially blood smears. Many alcohol mixtures may undergo slow reactions among ingredients upon long-term storage; in general most alcohol-based fixatives should be prepared no more than 1–2 days before use. Acetone fixation should be short (1 hour) at 4°C only on small specimens. Acetone produces extensive shrinkage and hardening, and results in microscopic distortion. It is used for immunohistochemistry, enzyme studies, and in the detection of rabies. Cold acetone is especially useful to ‘open’ the membranes of intact cells (e.g. grown on coverslips or microscope slides) to facilitate entrance of large molecules (e.g. antibodies for immunohistochemical studies). ‘Trade secret’ ingredients stabilize commercial formulations.

Clarke’s solution

Absolute ethanol 60 ml Glacial acetic acid 20 ml
This solution produces good general histological results for H&E stains. It has the advantage of preserving nucleic acids while lipids are extracted. A short fixation is recommended and tissues are transferred to 95% ethanol following fixation.

Carnoy’s fixative

Acetic acid 10 ml Absolute ethanol 60 ml Chloroform 30 ml
Carnoy’s fixative is useful for RNA stains, e.g. methyl green pyronine, and for glycogen preservation. It shrinks and hardens tissues and hemolyzes red blood cells. It may destroy the staining of acid-fast bacilli. It is useful in cytology to clear heavily blood-stained specimens.


Acetic acid 10 ml 100% methanol 60 ml Chloroform 30 ml
Causes less hardening and less shrinkage than Carnoy’s, but with the same pattern of staining.

Dehydrant cross-linking fixatives
Compound fixatives with both dehydrant and cross-linking actions include alcohol-formalin mixtures.
Alcohol-formalin fixation or post-fixation can be advantageous in large specimens with extensive fat. Lymph nodes can be detected much more easily in specimens with alcohol-formalin fixation due to the extraction of lipids and to texture differences compared with tissues fixed in NBF. The preparation of alcohol-formaldehyde solutions is complex, especially buffered forms of this compound fixative. It is probably best to purchase commercial preparations of buffered alcohol-formaldehyde. For use in post-fixation (e.g. after 10% NBF), Carson (1990) recommends the following formula:
Absolute ethanol 650 ml Distilled water 250 ml Formaldehyde (37%) 100 ml
Carson recommends this formula because she noted that the concentration of ethanol should be less than 70% to prevent the precipitation of phosphates in 10% NBF saturated tissues. For initial fixation the following formulae can be used:

Alcoholic formalin

Ethanol (95%) 895 ml Formaldehyde (37%) 105 ml

Alcohol-formalin-acetic acid fixative

Ethanol (95%) 85 ml Formaldehyde (37%) 10 ml Glacial acetic acid 5 ml
Methanol may be substituted for ethanol with care; similarly, various mixtures of ethanol, acetic acid, and formalin may be used.

Alcoholic Bouin’s (Gendre’s solution)
This fixative is similar to Bouin’s except it is less aqueous and there is better retention in tissues of some carbohydrates (e.g. glycogen). Fixation should be between 4 hours and overnight followed by washing in 70% ethanol, followed by 95% ethanol (several changes). This is the one alcoholic fixative that improves upon aging ( Lillie & Fullmer 1976 ).

Gendre’s solution

95% ethanol saturated with picric acid (5 g per 100 ml) 800 ml Formaldehyde (37%) 150 ml Glacial acetic acid 50 ml
To increase the effectiveness of alcoholic Bouin’s, if there is no time for aging, the following formula has been recommended ( Gregory 1980 ):

Equivalent to aged alcoholic Bouin’s

Picric acid 0.5 g Formaldehyde (37%) 15 ml 95% ethanol 25 ml Glacial acetic acid 5 ml Ethyl acetate 25 ml Tap water 30 ml

Another alcoholic form of Bouin’s solution is as follows:

Stock Bouin’s solution 75 ml 95% ethanol 25 ml
This solution is excellent for lymph nodes (24 hours) and for fatty tissue (48 hours).
A closely related fixative is:

Rossman’s solution

Tap water 10 ml Formaldehyde (37%) 10 ml Absolute ethanol 80 ml Lead nitrate 8 g
Fix for 24 hours at room temperature. This is a good fixative for connective tissue mucins and umbilical cord.

For metabolic bone disease

Phosphate buffer

Tap water 1000 ml NaH 2 PO 4 ·H 2 O 1.104 g NaHPO 4 (anhydrous) 4.675 g


Phosphate buffer 900 ml Formaldehyde (37%) 100 ml Adjust pH to 7.35.  

Fixation and decalcifation

Bouin’s decalcifying solution

Saturated aqueous solution of picric acid (10.5 g per 500 ml) 500 ml Formaldehyde (37%) 167 ml Formic acid 33 ml

Fixation for fatty tissue

Bouin’s solution 75 ml 95% ethanol 25 ml
May require up to 48 hours for good sections of lipomas or well-differentiated liposarcomas.

This chapter is an introduction to fixation. More detailed and advanced issues related to fixation are included in several other texts/references ( Sheehan & Hrapchak 1980 ; Eltoum et al. 2001a , 2001b ; Grizzle et al. 2001 ). As discussed, various formulae may vary within a few percentages, but most of these formulae produce equivalent results.


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5 The gross room/surgical cut-up

S. Kim Suvarna, Christopher Layton

The dissection and preparation of any specimen for histological/microscopic analysis involves more than simply the tissue processing and section cutting. Whilst the dissection and laboratory area are often perceived as the key elements of the department, it must be clearly understood that there are many other steps that follow specimen receipt. Some are specific to tissue selection and handling, and others are clearly support role in type. It is also implicit that a good laboratory will be adequately staffed by appropriately trained scientific/medical and support staff (secretarial, medical laboratory assistants, administration, etc.), as they interface at multiple levels with pathological sample handling. Indeed, a poorly staffed department will perform weakly – at best.

Safety first and last
The histopathology department is rich in hazards (e.g. biological, chemical, radiation). These are also risks reflecting the range of materials used to store, process and analyze tissues. These may be toxic, flammable, allergic, carcinogenic, electrical, etc. Furthermore, the presence of sharp cutting implements, complex machinery and the movement of the specimens around the laboratory needs staff to be fully trained and aware of all these potential hazards. Every laboratory should have accessible and clear standard operating procedures (SOPs, see Chapter 1 ), of which many will reflect national/international guidelines ( websites 1 – 3 ). Ongoing safety education is required and caution should be employed at every step of specimen handling for safe laboratory practice ( websites 2 and 4 ).

Specimen reception
A separate room is required for specimen reception, acting as the interface between hospital staff (or other visitors) and the pathological laboratory. Appropriate benching and good lighting must be available, along with good ventilation, safety equipment, disinfectants, absorption granules and protective clothing. In the event of specimen spillage (i.e., body fluids, fixative leakage or other mishap), the immediate response by staff will limit any potential local health risk, and also prevent risk to other laboratory personnel.
The key point of this room is to receive samples safely and securely. The specimen should be confirmed in terms of the identity and to be assigned a unique laboratory specimen identifier – usually a complex number. Correlation of the specimen against the clinical request form is mandatory, along with checking of appropriate clinical details mentioned against the specimen. Corroborative data in the form of the hospital number/registration index, national patient identifier number, the full name, date of birth and address are also valid ways of verifying the identity of any specimen. Multiple sources of cross reference are advocated, and if there is any doubt with regard to the probity of specimen then it should not be passed onwards until the clinician concerned has confirmed the appropriate details and probity of the sample.
In many situations the two-person rule is best followed, with two independent laboratory practitioners verifying the various details of the specimen – at all the different stages of examination. Confirmation of a minimum of three unique identifiers (as detailed above) is advisable. Once validated and identified, the case can be passed to the dissection room for examination, specimen description and block sampling.
The usual method of specimen identification is simply the year (expressed usually in two digits) with a sequential numbering system starting with one (1) and proceeding up to the final specimen of each year. There may be a check digit, usually in the form of a letter applied, but this simple system allows surgical pathology samples to be processed with ease and to be correlated against paraffin blocks, photographs and other tests (see below). Thus, case 2345 L/12 is the two thousand, three hundred and forty-fifth sample of the year 2012. The letter suffix (L) is a computer check datum to verify that the numerical data is valid.
Particular attention must be paid to cases with unusual names, or contrastingly, very common names. Names which have a variety of different spellings and any specimens that have incomplete information should be very carefully considered before being accepted.
In some cases multiple specimens from a single patient may be received on the same day for analysis. Some laboratories prefer to annotate each sample with a separate number. However, a single laboratory number may suffice, but with sub-parts of the specimen being separately designated (e.g., sample A, sample B, etc.). Within this framework, if multiple blocks were taken from a sub-part of the specimen then these can be designated with individual numbers/letters in a similar ascending fashion. Thus, a gastrectomy sample with lymph node groups and the spleen could have one case number, multiple sub-part specimens and multiple blocks that can be correlated against the surgeon’s operative dissection. For example, using the number described above, the spleen in this case could be being designated 2345 L/12.C.2 (C. indicating the sub-part of the third sample = spleen; and the block number = 2).
Barcodes can be used where appropriate facilities exist, but in general terms many laboratories still have paper request forms that will accompany the specimen as it passes through the laboratory and towards final report emission.

Surgical cut-up/specimen dissection/grossing
The ideal layout of this room is a matter of debate, varying between different laboratories and pathologists’ needs. There are multiple different design solutions existing around the general principles of a histology laboratory ( Rosai 2004 , Cook 2006 , websites 2 , 4 ), but it is imperative that the dissection area must have good lighting, good ventilation, non-absorbent wipe-clean surfaces, appropriate protective clothing for the laboratory personnel, gloves and other equipment (photography, tissue macerators, disposal bins). The dissection room should be a comfortable environment permitting undisturbed work by the pathologist and support technical staff. Given that the range of specimens received in most laboratories is wide, the technical staff will have to be familiar with the various requirements of different specimens that guide their subsequent handling and pathological preparation.
It is a matter of preference whether the operators within this environment sit or stand, and ideally both options should be available. Modern dissection areas often have integrated dissection desks, enclosed fluid/fixative feeds and laminar down-draft ventilation ( website 5 ) in order to protect both the dissector and support staff from formalin vapor. All tools and materials should be ergonomically accessible. The room should have good natural and/or electric lighting ( Fig. 5.1 ).

Figure 5.1 A pathology dissection station with a downdraft ventilated bench and clear dissection zone. Note the well-lit and ergonomic layout for the grossing pathologist and the technician support.
(Grateful thanks are expressed to Dr Caroline Verbeke and Mr Jonathan Sheriff for their assistance and consent for the illustration.)

Thinking before dissection
Prior to fixation it may be relevant to reserve some tissue from the specimens for microbiology assessment (being placed into appropriate culture media) and/or electron microscopy (requiring glutaraldehyde fixation). Fresh tissue can be taken for DNA extraction, cytogenetics and molecular pathology techniques. Other specialized tests (e.g., mass spectroscopy) may also require tissue retention before standard formalin fixation. Some samples need fixation and then decalcification in EDTA ( see section later in the book ).
Some specimens are only examined by means of macroscopic assessment, possibly with photography and other physical techniques. Examples include various mechanical/prosthetic implants, metal bodies, bullets, gallstones and medical devices. These must be dealt with according to the needs of the request/case. It should be noted that some specimens may require retention for a prolonged period of time – as in cases of forensic/criminal investigation.
This dissection/blocking/grossing/cut-up facility must have an appropriate storage area immediately to hand. This allows clearance of examined samples promptly, without the dissecting area becoming cluttered.
The individual choice of dissecting tools will reflect the type of specimen being considered ( Fig. 5.2 ). However, a range of cutting blades is advised, enabling the dissector to deal with small specimens through to complex and large resections. Very large knives are particularly useful for obtaining full transverse sections of organs (lungs, liver, etc.). The smaller blades are useful for precise trimming of tissues. However, before any knife is put to the specimen, it is emphasized that the tissue specimen should be well fixed. Forceps and absorbent cloths should be available. The blocks taken (vi) should not completely fill the cassette ( Fig. 5.3 ) as this would impede processing fluid access to the tissue. Thus, tissue cassettes are generally made of plastic and conform to a variety of size standards across the developed world nowadays. Most standard blocks allow a sample of about 20 × 20 × 3 mm thick tissue to be contained and processed. There is variation in cassette size that does allow larger blocks to be selected ( Fig. 5.2 ). This is particularly useful for histological examination of large surgical resections where the global geography of the specimen is needed for analysis. Examples could include rectal cancer resection, radical prostatectomy and autopsy lung tissue for industrial disease. However, some general rules can be developed to specimen handling/sampling.

Figure 5.2 Cut-up/grossing tools. A range of small and large bladed tools are advocated along with forceps, ruler and a fluid-resistant dissecting surface. An appropriate measure and access to photography are needed. Varying sizes of cassette (centrally) are available, in a range of colors and sizes, to permit handling of varying amounts of sample and also to indicate handling issues that follow tissue processing.

Figure 5.3 Tissue blocks are placed into the cassette. Note they should not fill the cassette, and must permit room for processing fluid circulation. The orientation of the blocks is enhanced by a sponge securing the specimens in sequential position and a colored agar marker allows designation of the order of slices taken. The samples have been marked with different colored inks to permit designation of the sidedness of the samples and the resection margins.
The specimens should be analyzed with only one pot open at any one time. The request and specimen identity should be checked. The sample should be described in terms of the shape, size and defining characteristics of the specimen. This means that small biopsies, for example endoscopic mucosal samples, may simply be afforded a simple descriptor in the form of the number of pieces and the size (SI units, usually mm) of the largest piece of tissue. An example could be ‘three pieces of brown tissue, the largest 3 mm diameter’.
Medium and large specimens ( Fig. 5.4 ) need more detailed and careful description of the various anatomical components, together with identification of macroscopic landmarks, orientation markers/sutures and the lesion/s as relevant. The background tissues, beyond the lesion under consideration, also require description. The sampling of any large case/resection should follow local/national guidelines in order to provide the relevant information for subsequent clinical management of the patient ( website 6 ).

Figure 5.4 A medium-sized skin sample is seen with a central lesion. This could be described as ‘A skin ellipse x by y by z mm depth is seen with an orientation suture, designated 12 o’clock. The sample shows a central yellow-brown nodule z mm that is k mm clear of the closest margin’. It is sectioned into parallel slices and then placed into a cassette (see Fig. 5.3 ).
Any macroscopic description is usually dictated for subsequent secretarial transcription, or on occasion can be simply written down for typing later. Canned/proforma reports may be of value to standardize the approach to samples. The departmental computer system can be set to track specimen movement through the laboratory, from receipt to final pathologist report authorization. Photographing the macroscopic specimen is particularly important in cases of complex surgical excision (e.g. Wertheim’s hysterectomy, pneumonectomy, AP resection, etc.), and may be of use in later analysis/case discussion ( Fig. 5.5 ). The availability of cheap and reliable digital photography has been a major bonus to the laboratory, permitting verification of the image being captured. Nevertheless, specialist photography may still be required for cases that might end up as visual teaching presentation, journal/book illustration publication, or in a medico-legal setting. Consideration of the potential to recognize a patient’s sample should be made, with some guidance existing on the subject ( website 7 ).

Figure 5.5 A lung lobectomy sample, sliced to show the hilar and mediastinal plane of resection, highlighting tumor adjacent to the margin. Block sampling at this interface and background tissue sampling against standard protocols will allow full analysis. Note the numbered cassette along with the ruler for full case identification and analysis.
If one knows beforehand of additional tests that are automatically required on some specimens (such as liver core biopsies requiring multiple ancillary histochemical stains) then different color cassettes/ markers can be used in order to designate additional actions that should follow as an automatic laboratory consequence ( Fig. 5.2 ). The different colored cassettes may also indicate the types of section and sections required, as well as the speed/urgency of any specimen.
Following dissection, the residual tissues must be stored in a ventilated secure archive format, and waste materials must be disposed of according to local health and safety regulations ( websites 4 , 5 , 8 ).

Specimen dissection plans

Small samples
Small biopsy samples rarely need dissection, and can be simply processed, embedded and then sectioned as they present themselves. In some cases orientation of the specimen can be facilitated by means of a dissecting microscope or magnifying lens – such as when considering morphological abnormalities of small bowel biopsies. However, the majority of small biopsies can be adequately examined at multiple levels allowing the pathologist to mentally reconstruct the three-dimensional quality of the tissue during microscopic examination.
These small samples may benefit from being placed in a nylon bag, between metal disks with fine mesh, within paper, etc., in order to prevent them falling through the cassette perforations and being lost. In some cases, eosin is used as a marker for small samples, in order to highlight them on the background of paper, embedding bench or equivalent. It is recommended that a count of the small tissue biopsy fragments is taken at the description/ grossing stage in order to verify that all the tissue has survived processing prior to section cutting.

Core biopsies
These are treated in a somewhat similar manner, although their embedding requires being laid out in longitudinal fashion so that the plane of section cuts along the majority of the tissue. Larger cores (with diameters of 4–5 mm or greater) may occasionally benefit by division into two halves along the long axis. Alternatively it may be easier to simply provide multiple levels with retention of tissue in between the levels, for adequate analysis. Multiple cores often require each core being placed into individual cassettes.

Skin biopsies
These include simple punch biopsies (handled akin to cores) and shave biopsies that should be mounted on edge in order to provide an adequate view of the epidermis, dermis and subcuticular substrates. A marker item placed into the cassette (e.g., plastic bead or colored paper) will identify such samples to embedding personnel. Alternatively, some laboratories use cheese paste to help maintain specimen orientation ( Tripathi et al. 2008 ). The protein in the paste helps hold the tissue orientation during processing ( Fig. 5.6 ).

Figure 5.6 Cheese paste is seen holding the thin fragment of inked skin on edge and in position securely. The cheese protein matrix will survive tissue processing!
Skin samples also include the more complex intermediate and large specimens for removal of defined lesions, right up to radical skin cancer resections including deep soft/bony tissues.
The intermediate and larger samples of skin are often presented as an ovoid/ellipse/piece of skin/subcutis, mostly with a central lesion (see Fig. 5.4 ). These must be described in terms of the width and breadth of the specimen together with a depth. The lesion characteristics (nodule, ulcer, papule, color/margin, etc.) should be discussed. Specimens of these skin resections are often best managed in sequential/serial transverse section, with Indian ink/other dyes being applied to different surfaces in order to confirm the orientation/boundaries of the specimen ( Fig 5.3 ). Markers can be placed into cassettes in order to confirm pieces of tissue with orientation markers, although it is generally found that specimens start with small apical transverse sections through to the broadest point across the waist of the specimen and then taper off towards the other end.
Very large resections of skin with soft tissues may require photography and then targeted block sampling. This should also allow for the appropriate assessment of any tumor/lesion with deep and lateral margin correlation along with multiple blocks of the pathology in order to allow for disease variations that may be present. These samples, and indeed their smaller counterparts, should be blocked to permit reporting against national/international standards.

Bowel specimens
These generally are medium and large tissue resections along the length of the gastrointestinal tract (e.g. partial colectomy/gastrectomy). They are best sampled with multiple (usually n ≥ 3) blocks of any lesion in relation to the adjacent mucosa, wall and serosal aspect tissues ( Fig. 5.7 ), although large geographic blocks can be employed. The margins often need inking and the background tissues including resection margins are often included as part of the relevant dissection protocol ( Allen 2006 , Allen & Cameron 2004 , website 6 ). Particular attention is paid to the lymph nodes, and these can be either manually dissected in groups, or can be identified from fat-clearance protocols ( Prabhudesai et al. 2005 ) as described below. It is vital that the nodes are assessed in terms of their proximity to the lesion along with their different ‘level’ stages. Many cases require consideration of the high tie (i.e. the most proximal node in the resection) lymph node (or equivalent).

Figure 5.7 A large bowel specimen is seen with anatomical complexity requiring a good description and multiple blocks to be taken for full analysis. This sample shows the resection, opened to visualize the cancer. The specimen has been photographed to facilitate understanding of the local resection margins and the serosal surface (inked). The entire tumor and local bowel can be blocked into a large cassette, if desired. A lymph node (arrowed) is clearly involved by tumor in the fatty serosal tissues, but the fatty mesentery can be removed for fat-clearance nodal identification and analysis.

Fat clearance
Finding lymph nodes within a large amount of fatty mesenteric/soft tissue can be problematic, and the ability to remove the adipose substrate from any specimen will lead to an enhanced rate of node detection and thereby sampling. One aspect of the histological tissue handling in the cut-up room allows such node identification ( Prabhudesai et al. 2005 ). The fatty tissue is usually sliced into 10 mm fragments and placed into large cassettes, thereby increasing the access of solvents to the specimen. Fat removal occurs as part of the processing of tissues, but the blocks of fatty parenchyma are normally removed from the processing chamber before the tissues are impregnated finally with wax. At this stage the lymph nodes can now be readily identified by transillumination of the tissue sample from below ( Fig. 5.8 ). The sampled nodes can then be placed back in the tissue processor in a smaller cassette, with normal embedding, sectioning and staining to follow.

Figure 5.8 Following fat clearance, the transilluminated sample is searched for nodes (arrowed). These are then extracted and placed into smaller cassettes for routine histology assessment after the final stage of processing and embedding.

Lung tissues
These generally are performed as localized biopsies or lobectomy/pneumonectomy specimens. The background pleura and lung must be evaluated along with any lesions as required in standard proforma sampling protocols ( Allen 2006 , Allen & Cameron 2004 , website 6 ). In general terms, multiple blocks for any tumor (usually n ≥ 3) along with sampling of pleural/mediastinal/bronchial margins are needed. Nodes are often presented in groups separately, although careful dissection of the hilar tissues should allow further node harvest from these tissues ( Fig. 5.5 ).

Gynecological samples
Common samples, such as cone biopsies from the cervix, need appropriate inking of margins and orientation, often in a serial block fashion across the specimen with specimen photography. This allows the three-dimensional assessment of dysplasia or invasive neoplasia in relation to the various surgical margins. Uterine samples are usually sampled in terms of background cervix, endometrium and myometrial tissues together with some representative sampling of common benign lesions (fibroids/leiomyomas/etc.). Sampling will be guided by local practice together with national guidelines ( Allen 2006 , Allen & Cameron 2004 , website 6 ). Dysplastic and malignant lesions often require multiple blocks, resection margins together with careful examination of related lymph nodes (usually presented, and therefore blocked, separately). Specific tissues such as tubes and ovaries should follow similar standard guidelines in terms of the sampling pattern, number of blocks and related tissue samples. It is emphasized that pluripotential differentiation of tumors within the female genital tract requires multiple blocks of a tumor to be taken for full analysis.

Breast resections
These are also common resections, usually with the need for inked margins, in relation to the orientation of the specimen. Multiple blocks of the tumor are usually required. Background tissues at multiple points should be also assessed and the lymph nodes (if present) are often examined in tiered/grouped fashion in order to assess tumor spread up towards the highest-level nodal size. Fat clearance may be required to capture all the nodes in the axillary tail.

Soft tissue resections
These should be examined with multiples blocks of tissue, background parenchyma and the margins. Some experts advocate 1 block for every 10 mm diameter of tumor, up to 10 blocks – although more may be required on occasion. Careful slicing and examination of the specimens macroscopically will allow sampling and consideration of all peripheral boundaries. Furthermore, given the pervasive nature of soft tissue tumors, this widespread sampling is usually required. Tumor sampling before fixation for molecular/genetic analysis may be required.

Other samples
The chapter is not sufficient to discuss all resections and specimen subtypes, and the groups above are illustrative only. The reader is referred to relevant governing bodies/organizations that have produced protocols for the analysis of other specimens ( Allen 2006 , Allen & Cameron 2004 , website 6 ).


Allen D.C. Histopathology reporting. Guidelines for surgical cancer , second ed. Springer, London; 2006.
Allen D.C., Cameron R.I. Histopathology specimens: clinical, pathological and laboratory aspects. London.: Springer, 2004.
Cook, D.J., 2006. Some aspects of the organisation of a histology laboratory. In: Cellular pathology, second ed. Scion, Bloxham, Oxfordshire, pp. 357–367.
Prabhudesai A.G., Dalton R., Kumar D., Finlayson A.G. Mechanised one-day fat clearance method to increase the lymph node yield in rectal cancer specimens. British Journal of Biomedical Science . 2005;62:120–123.
Rosai J. Introduction and gross techniques in surgical pathology. Rosai and Ackerman’s surgical pathology , ninth ed. Edinburgh: Mosby; 2004. pp. 1–24, 25–36
Tripathi M., Sethuraman C., Lindley R., Ali R.B. Comparison of use of cream cheese and agar gel for orientation of skin biopsies. XXIX Symposium of the ISDP, Graz, Austria, October 2–4, 2008. American Journal of Dermatopathology . 2008;30:514–533.


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6 Tissue processing

Lena T. Spencer, John D. Bancroft


Wanda Grace Jones

After the removal of a tissue sample from the patient, a series of physical and chemical processes must take place to ensure that the final microscopic slides produced are of a diagnostic quality. Tissues are exposed to a series of reagents that fix, dehydrate, clear, and infiltrate the tissue. The tissue is finally embedded in a medium that provides support for microtomy. The quality of the structural preservation of tissue components is determined by the choice of exposure times to the reagents during processing. Every step in tissue processing is important; from selection of the sample, determining the appropriate protocols and reagents to use, to staining and final diagnosis. Producing quality slides for diagnosis requires skills that are developed through continued practice and experience. As new technology and instrumentation develops, the role of the histology laboratory in patient care will continue to evolve, providing standardization of processes, increased productivity, and better utilization of the resources available. This chapter will provide an overview of the steps in the process and the reagents needed to prepare tissue for microscopic evaluation.

Labeling of tissues
A unique accession number or code should be assigned to every tissue sample as discussed in Chapter 5 . This unique number should accompany the specimens throughout the entire laboratory process and may be electronically or manually generated. New technology has made bar code quick response (QR) and character recognition systems readily available in most laboratories. Automated pre-labeling systems that permanently etch or emboss tissue cassettes and slides, as well as chemically resistant pens, pencils, slides and labels, are routinely used in pathology laboratories. Regardless of whether an automated or manual labeling system is used, adequate policies and procedures must be in place to ensure positive identification of the tissue blocks and slides during processing, diagnosis, and filing.

Principles of tissue processing
Tissue processing is designed to remove all extractable water from the tissue, replacing it with a support medium that provides sufficient rigidity to enable sectioning of the tissue without parenchymal damage or distortion.

Factors influencing the rate of processing
When tissue is immersed in fluid, an interchange occurs between the fluid within the tissue and the surrounding fluid. The rate of fluid exchange is dependent upon the exposed surface of the tissue that is in contact with the processing reagents. Several factors influence the rate at which the interchange occurs: namely, agitation, heat, viscosity and vacuum.

Agitation increases the flow of fresh solutions around the tissue. Automated processors incorporate vertical or rotary oscillation, or pressurized removal and replacement of fluids at timed intervals as the mechanism for agitation. Efficient agitation may reduce the overall processing time by up to 30%.

Heat increases the rate of penetration and fluid exchange. Heat must be used sparingly to reduce the possibility of shrinkage, hardening or embrittlement of the tissue sample. Temperatures limited to 45°C can be used, but higher temperatures may be deleterious to subsequent immunohistochemistry.

Viscosity is the property of resistance to the flow of a fluid. The smaller size of the molecules in the solution, the faster the rate of fluid penetration (low viscosity). Conversely, if the molecule size is larger, the rate of exchange is slower (high viscosity). Most of the solutions used in processing, dehydration and clearing, have similar viscosities, with the exception of cedar wood oil. Embedding mediums have varying viscosities. Paraffin has a lower viscosity in the fluid (melted) state, enhancing the rapidity of the impregnation.

Using pressure to increase the rate of infiltration decreases the time necessary to complete each step in the processing of tissue samples. Vacuum will remove reagents from the tissue, but only if they are more volatile than the reagent being replaced. Vacuum used on the automated processor should not exceed 50.79 kPa to prevent damage and deterioration to the tissue. Vacuum can also aid in the removal of trapped air in porous tissue. Impregnation time for dense, fatty tissue can be greatly reduced with the addition of vacuum during processing.

Stages of tissue processing

• Fixation – stabilizes and hardens tissue with minimal distortion of cells.
• Dehydration – removal of water and fixative from the tissue.
• Clearing – removal of dehydrating solutions, making the tissue components receptive to the infiltrating medium.
• Infiltrating – permeating the tissue with a support medium.
• Embedding – orienting the tissue sample in a support medium and allowing it to solidify.

Preserving cells and tissue components with minimal distortion is the most important aim of processing tissue samples. Fixation stabilizes proteins, rendering the cell and its components resistant to further autolysis by inactivating lysosomal enzymes. It also changes the tissues’ receptiveness to further processing. Fixation must finish before subsequent steps in the processing schedule are initiated.
If fixation is not complete prior to processing, stations should be designated on the processor for this purpose. If the tissue is inadequately fixed, the subsequent dehydration solutions may complete the process, possibly altering the staining characteristics of the tissue. The size and type of specimen in the tissue cassette determines the time needed for complete fixation and processing. The tissue should be dissected to 2–4 mm in thickness. Care must be taken not to overfill the cassette, as this would impede the flow of reagents around the tissue. If possible, larger and smaller pieces of tissue should be separated and processed using different schedules. The most commonly used reagent for the fixation of histological specimens is 10% neutral buffered formalin (NBF) – see Chapter 4 .

Post-fixation treatment
Special fixation techniques may require additional steps before processing is initiated. Picric acid fixatives (Bouin’s) form water-soluble picrates making it necessary to place the tissue cassettes directly into 70% alcohol for processing. Alcoholic fixatives, such as Carnoy’s fluid, should be placed directly into 100% alcohol. To help in the visualization of small fragments of tissue during embedding, a few drops of 1% eosin can be added to the specimen container 30 minutes prior to processing. The pink color of the tissue remains during processing, but washes out during subsequent staining.

The first stage of processing is the removal of ‘free’ unbound water and aqueous fixatives from the tissue components. Many dehydrating reagents are hydrophilic (‘water loving’), possessing strong polar groups that interact with the water molecules in the tissue by hydrogen bonding. Other reagents affect dehydration by repeated dilution of the aqueous tissue fluids. Dehydration should be accomplished slowly. If the concentration gradient is excessive, diffusion currents across the cell membranes may increase the possibility of cell distortion. For this reason, specimens are processed through a graded series of reagents of increasing concentration. Excessive dehydration may cause the tissue to become hard, brittle and shrunken. Incomplete dehydration will impair the penetration of the clearing reagents into the tissue, leaving the specimen soft and non-receptive to infiltration. There are numerous dehydrating agent; ethanol, ethanol acetone, methanol, isopropyl, glycol and denatured alcohols.

Dehydrating fluids

Ethanol (C 2 H 5 OH)
Ethanol is a clear, colorless, flammable liquid. It is hydrophilic, miscible with water and other organic solvents, fast-acting and reliable. Aside from its human health-risk potential, ethanol is taxable, controlled by many governments, and therefore requires careful record keeping. Graded concentrations of ethanol are used for dehydration; the tissue is immersed in 70% ethanol in water, followed by 95% and 100% solutions. Ethanol ensures total dehydration, making it the reagent of choice for the processing of electron microscopy specimens. For delicate tissue it is recommended that the processing starts in 30% ethanol.

Industrial methylated spirit (denatured alcohol)
This fluid has the same physical property as ethanol. Denatured alcohol consists of ethanol, with the addition of methanol (about 1%), isopropyl alcohol or a combination of alcohols. For purposes of tissue processing it is used in the same manner as ethanol.

Methanol (CH 3 OH)
Methanol is a clear, colorless and flammable fluid that is miscible with water, ethanol and most organic solvents. It is highly toxic but can be substituted for ethanol in processing protocols.

Propan-2-ol, isopropyl alcohol (CH 3 CHOHCH 3 )
Isopropyl alcohol is miscible with water, ethanol and most organic solvents. It is used in microwave processing schedules. Isopropyl alcohol does not cause over-hardening or shrinkage of the tissue.

Butyl alcohol (butanol) (C 4 H 9 OH)
Used primarily for plant and animal histology. Butyl alcohol is a slow dehydrant causing less shrinkage and hardening of the tissue.

Acetone (CH 3 COCH 3 )
Acetone is a clear, colorless, flammable fluid that is miscible with water, ethanol and most organic solvents. It is rapid in action, but has poor penetration and causes brittleness in tissues if its use is prolonged. Acetone removes lipids from tissue during processing.

Additives to dehydrating agents
When added to dehydrating agents, phenol acts as a softening agent for hard tissues such as tendon, nail, and dense fibrous tissue and keratin masses. Phenol (4%) should be added to each of the 95% ethanol stations. Alternatively, hard tissue can be immersed in a glycerol/alcohol mixture.

Universal solvents
Universal solvents are no longer used for routine processing due to their hazardous properties, and they should be handled with extreme care. Universal solvents both dehydrate and clear tissues during tissue processing. Dioxane, tertiary butanol and tetrahydrofuran are considered to be universal solvents. They are not recommended for processing delicate tissues due to their hardening properties.

Clearing reagents act as an intermediary between the dehydration and infiltration solutions. They should be miscible with both solutions. Most clearants are hydrocarbons with refractive indices similar to protein. When the dehydrating agent has been entirely replaced by most of these solvents the tissue has a translucent appearance: hence the term ‘clearing agent’.
The criteria for choosing a suitable clearing agent are:

• rapid penetration of tissues
• rapid removal of dehydrating agent
• ease of removal by melted paraffin wax
• minimal tissue damage
• low flammability
• low toxicity
• low cost
Most clearing agents are flammable liquids, which warrant caution in their use. The boiling point of the clearing agent gives an indication of its speed of replacement by melted paraffin wax. Fluids with a low boiling point are generally more readily replaced. Viscosity influences the speed of penetration of the clearing agent. Prolonged exposure to most clearing agents causes the tissue to become brittle. The time in the clearing agent should be closely monitored to ensure that dense tissue blocks are sufficiently cleared and smaller more fragile tissue blocks are not damaged. Cost should be considered, especially as it relates to disposal of the reagent. Since most clearing agents are aromatic hydrocarbons or short-chain aliphatic hydrocarbons, environmental issues must be addressed. Most institutions have a policy for the storage, disposal and safety requirements for all flammables used in the laboratory.

Clearing agents suitable for routine use

Xylene is a flammable, colorless liquid with a characteristic petroleum or aromatic odor, which is miscible with most organic solvents and paraffin wax. It is suitable for clearing blocks that are less than 5 mm in thickness and rapidly replaces alcohol from the tissue. Overexposure to xylene during processing can cause hardening of tissues. It is most commonly used in routine histology laboratories and is also recyclable.

This has similar properties to xylene, although it is less damaging with prolonged immersion of tissue. It is more flammable and volatile than xylene.

Chloroform is slower in action than xylene but causes less brittleness. Thicker tissue blocks can be processed, greater than 1 mm in thickness. Tissues placed in chloroform do not become translucent. It is non-flammable but highly toxic, and produces highly toxic phosgene gas when heated. It is most commonly used when processing specimens of the central nervous system.

Xylene substitutes
Xylene substitutes are aliphatic hydrocarbons that exist in long- and short-chained forms. They differ in the number of carbon atoms within the carbon chain. Short-chained aliphatics have the same evaporation properties as xylene, and have no affinity for water. Long-chained aliphatics do not evaporate rapidly and may cause contamination of the paraffin wax on tissue processors.

Citrus fruit oils – limonene reagents
Limonene reagents are extracts from orange and lemon rinds; they are non-toxic and miscible with water. Disposal is dependent upon the water treatment centers and local/national standards. The main disadvantages are that they can cause sensitization and have a strong pungent odor that may cause headaches. Also, small mineral deposits such as copper or calcium may dissolve and leach from tissues. They are extremely oily and cannot be recycled.

Infiltrating and embedding reagents

Paraffin wax
Paraffin wax continues to be the most popular infiltration and embedding medium in histopathology laboratories. Paraffin wax is a mixture of long-chained hydrocarbons produced in the cracking of mineral oil. Its properties are varied depending on the melting point used, ranging from 47 to 64°C. Paraffin wax permeates the tissue in liquid form and solidifies rapidly when cooled. The tissue is impregnated with the medium, forming a matrix and preventing distortion of the tissue structure during microtomy. It has a wide range of melting points, which is important for use in the different climatic regions of the world. To promote desirable ribboning during microtomy, paraffin wax of suitable hardness at room temperature should be chosen. Heating the paraffin wax to a high temperature alters the properties of the wax. Higher melting point paraffin wax provides better support for harder tissues, e.g. bone, can allow production of thinner sections, but may cause difficulty with ribboning. Lower melting point paraffin wax is softer and provides less support for harder tissues. It is more difficult to obtain thinner sections but ribboning is easier. Paraffin wax is inexpensive, provides quality sections and is easily adaptable to a variety of uses. Paraffin wax is compatible with most routine and special stains, as well as immunohistochemistry protocols.

Paraffin wax additives
Paraffin waxes that contain plasticizers or other resin additives are commercially available, providing a selection that is appropriate for most laboratories. These additives create paraffin waxes with selectable hardness compatible with the tissue to be embedded. The amount of additive will impact the rate of infiltration. Substances added to paraffin wax in the past include beeswax, rubber, ceresin, plastic polymers and diethylene glycol distearate. Many of these additives had a higher melting point than paraffin wax, consequently making the tissue more brittle.

Alternative embedding media
There are occasions when paraffin wax is an unsuitable medium for the type of tissue being processed including:

• Processing reagents remove or destroy tissue components that are the object of investigation, e.g. lipids
• Sections are required to be thinner, e.g. lymph nodes
• The use of heat may adversely affect tissues or enzymes
• The infiltrating medium is not sufficiently hard to support the tissue

Resin is used exclusively as the embedding medium for electron microscopy (see Chapter 22 ), ultra-thin sectioning for high resolution and also for undecalcified bone (see Chapter 16 ).

Agar gel alone does not provide sufficient support for sectioning tissues. Its main use is as a cohesive agent for small friable pieces of tissue after fixation, a process known as double embedding. Fragments of tissue are embedded in melted agar, allowed to solidify and trimmed for routine processing. A superior, more refined, method is to filter the fixative containing small, friable tissue fragments through a Millipore filter using suction. Molten agar is then carefully poured into the filter apparatus, the agar is left to solidify and the resultant agar pellet is removed and routinely processed and embedded in paraffin wax.

Gelatin is primarily used in the production of sections of whole organs using the Gough-Wentworth technique and in frozen sectioning. It is rarely used.

The use of celloidin or LVN (low viscosity nitrocellulose) is discouraged because of the special requirements needed to house the processing reagents and the limited use these types of sections have in neuropathology. It is rarely used.

Paraffin wax embedding
Embedding involves the enclosing of properly processed, correctly oriented specimens in a support medium that provides external support during microtomy. The embedding media must fill the matrix within the tissue, supporting cellular components. The medium should provide elasticity, resisting section distortion while facilitating sectioning.
Most laboratories use modular embedding centers, consisting of a paraffin dispenser, a cold plate, and a heated storage area for molds and tissue cassettes. Paraffin wax is dispensed automatically from a nozzle into a suitably sized mold. The tissue is oriented in the mold; a cassette is attached, producing a flat block face with parallel sides. The mold is placed on a small cooling area to allow the paraffin wax to solidify. The quick cooling of the wax ensures a small crystalline structure, producing fewer artifacts when sectioning the tissue.

Orientation of tissues
Specimen orientation during embedding is important for the demonstration of proper morphology. Incorrect orientation may result in diagnostic tissue elements being damaged during microscopy or not being evident for pathology review. Products are available that help ensure proper orientation: marking systems, tattoo dyes, biopsy bags, sponges, and papers. Orientation of the tissue should offer the least resistance of the tissue against the knife during sectioning. A margin of embedding medium around the tissue assures support of the tissue.
Tissues requiring special orientation include:

• Tubular structures: cross section of the wall and lumen should be visible; arteries, veins, fallopian tube and vas deferens samples.
• Skin biopsies; shave punch or excisions, cross section of the epidermis, dermis and subcutaneous layers must be visible.
• Intestine, gallbladder, and other epithelial biopsies: cut in a plane at right angles to the surface, and oriented so the epithelial surface is cut last, minimizing compression and distortion of the epithelial layer.
• Muscle biopsies: sections containing both transverse and longitudinal planes.
• Multiple pieces of a tissue are oriented side by side with the epithelial surface facing in the same direction.

Automated tissue processing
The basic principle for tissue processing requires the exchange of fluids using a series of solutions for a predetermined length of time in a controlled environment. For decades, the instrumentation used in tissue processing remained relatively unchanged. Recent advances now include specialty microwave ovens, the emergence of constant throughput processors, and processors with multi-sectioned retorts.

Tissue processors
The carousel-type processor (tissue transfer) and the self-contained fluid exchange systems were the first automated tissue processors used in the histology laboratory. The carousel-type processor transports tissue blocks contained in baskets through a series of reagents housed in stationary containers. The length of time the specimens were submerged in each reagent container was electronically controlled. Earlier models accomplished this step by notching the face of a clock disk. Vertical oscillation or the mechanically raising and lowering of the tissue into the reagent containers provided the agitation needed for the processing of the tissue.
The enclosed, self-contained vacuum tissue processor later became the mainstay of most laboratories. A microprocessor was used to program this instrument. Tissues were loaded into a retort chamber where they remained throughout the process. Reagents and melted paraffin wax were moved sequentially into and out of the retort chamber using vacuum and pressure. Each step could be customized by controlling time, temperature, or pressure/vacuum (P/V). The advantages of this system are that vacuum and heat can be used at any stage, customized schedules for tissue processing are possible, and there is fluid spillage containment and elimination of fumes. These processors usually employ alarm systems and diagnostic programs for troubleshooting any instrumentation malfunction. Newer instrumentation have divided retorts that allows for different programs to run simultaneously, allowing for better utilization of the equipment and providing the opportunity to divide tissue by size. Many processors have solution management systems, allowing reagents to be monitored for purity and to be used for a greater period of time without adversely harming the tissue.

Microwave processors
Microwave ovens specially designed for tissue processing are now common. The microwave oven shortens the processing time from hours to minutes. Microwave exposure stimulates the diffusion of the solutions into the tissue by increasing the internal heat of the specimen, thus accelerating the reaction. Tissues are manually transferred from container to container of reagent. Most laboratory microwave ovens contain precise temperature controls, timers, and fume extraction systems. The processing time depends on the thickness and density of the specimen. Reagents used for microwave processing include ethanol, isopropanol and proprietary mixtures of alcohol, and paraffin. Graded concentration of solutions is not required. Clearing agents are not necessary because the temperature of the final paraffin step facilitates evaporation of the alcohols from the tissue. Xylene and formalin are not used in this process, which eliminates toxic fumes and carcinogens. Properly controlled processing provides uncompromised morphology and antigenicity of the specimens. Increased efficiency through improved turnaround times, environmentally friendly reagents, and greater profitability due to reduction in number and volume of reagents are advantages of this system. Disadvantages of the system include the fact that the process is labor intensive because the solutions are manually manipulated, temperatures must be maintained between 70 and 85°C, and the size of tissue sample is critical (2 mm). Also the cost of laboratory-grade microwaves may be prohibitive, and proper use of the microwave oven requires careful calibration and monitoring.

Alternative rapid processors
Advances in technology have led to the development of a ‘continuous input rapid tissue processor’. The enclosed processor uses microwave technology, vacuum infiltration and proprietary reagents which are described as being ‘molecular-friendly’. A robotic arm moves the tissue cassettes through four stations which contain acetone, isopropanol, polyethylene glycol, mineral oil and paraffin. Microwaves and agitation are used to accelerate the diffusion of solvents in tissue. A patented microwave technology is utilized, which operates at a continuous low power instead of pulsing high levels of microwave energy. The retort chamber is cylindrical; microwaves circle around the cavity, taking advantage of the physical principle of the ‘whispering chamber’ effect that eliminates hot and cold spots. The advantage of this system is the acceptance of tissues into the system every 15 minutes, improving turnaround time. The reagents used are environmentally safe, eliminating toxic vapors in the laboratory. The morphology and quality of the specimens is consistent with that of traditional tissue processing. The disadvantages include the cost of the processor, and the grossing of the tissue sample that requires standardization of specimen dissection.

Advantages of newer technology in processing

• custom programs specific to tissues being processed, addition of vacuum, agitation or heat at any stage
• rapid schedules
• fluid and fume containment
• environmentally friendly reagents
• time delay for start of processing schedules
• reagent management

Processor maintenance
Every institution should have a policy outlining the rotation and changing of solutions on the tissue processor. The numbers, sizes, types of tissue processed and the reagents used will play a role in the determination of this policy. Solutions should be carefully monitored to ensure quality. Every manufacturer has a handbook outlining an appropriate maintenance schedule.

Important maintenance tips

• Any spillage or overflow should be cleaned immediately
• Accumulation of wax on any surface should be removed
• The temperature of the paraffin wax bath should be set to 3°C above the melting point of the paraffin wax and monitored daily
• Timings should be checked when placing tissue cassettes in the processor, especially when delayed schedules are selected
• Warm water flushes should be incorporated, keeping the lines free of salts, protein and debris

Automated processing schedules
Although overnight schedules for tissue processing remain popular in many laboratories, schedules have changed to reflect the emphasis on reducing turnaround time for specimen reporting. Rapid processing for small biopsies or stat specimens can easily be accommodated.

Overnight processing
For many laboratories, this is considered to be the routine processing schedule. Tissues continue fixation by being immersed in 10% formalin, buffered or unbuffered. The process may include alcoholic formalin, varying concentrations of alcohol, xylene, or a xylene substitute, followed by infiltration in paraffin wax. Schedules are customized for the tissues being processed. Factors influencing the processing schedule include end time required, reagents used, the inclusion of heat and vacuum and the size and number of tissue cassettes processed. The schedule in Table 6.1 can be modified, adjusting times for the various stations, keeping in mind the end time needed for process completion.

Table 6.1 Overnight processing schedule

Processing breast specimens
Standardization of the fixation and processing of breast tissue has become a focus of laboratory regulatory agencies. Pre-analytic, analytic and post-analytic variables are addressed in the findings. The purpose of the guidelines is to improve the accuracy of hormone receptor testing and the utility of these prognostic and predictive markers for assessing breast carcinomas.
Due to the pressures of compliance with the new standards and guidelines, histology laboratories are revisiting their processing schedules, and adapting or adjusting new protocols for breast tissue. Currently, breast tissue should be received in the histology laboratory within one hour of removal from the patient, sectioned at 5 mm intervals and placed in 10% neutral buffered formalin and fixed for no less than six hours and not more than 72 hours before processing. Table 6.2 provides a processing schedule that is adjustable to address the parameters of these guidelines.

Table 6.

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